Title: <i>In vivo</i> generation of a mature and functional artificial skeletal muscle
Abstract: Research Article25 February 2015Open Access Source Data In vivo generation of a mature and functional artificial skeletal muscle Claudia Fuoco Claudia Fuoco Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Roberto Rizzi Roberto Rizzi IRCCS MultiMedica, Milan, Italy Cell Biology and Neurobiology Institute, National Research Council of Italy, Rome, Italy Search for more papers by this author Antonella Biondo Antonella Biondo Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Emanuela Longa Emanuela Longa Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy Search for more papers by this author Anna Mascaro Anna Mascaro Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy Search for more papers by this author Keren Shapira-Schweitzer Keren Shapira-Schweitzer Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel Search for more papers by this author Olga Kossovar Olga Kossovar Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel Search for more papers by this author Sara Benedetti Sara Benedetti Department of Cell and Developmental Biology, University College London, London, UKCorrection added on 10 March 2015, after first online publication: author affiliations have been corrected. Search for more papers by this author Maria L Salvatori Maria L Salvatori Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Sabrina Santoleri Sabrina Santoleri Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Stefano Testa Stefano Testa Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Sergio Bernardini Sergio Bernardini Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Roberto Bottinelli Roberto Bottinelli Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy Fondazione Salvatore Maugeri (IRCCS), Scientific Institute of Pavia, Pavia, Italy Search for more papers by this author Claudia Bearzi Claudia Bearzi IRCCS MultiMedica, Milan, Italy Cell Biology and Neurobiology Institute, National Research Council of Italy, Rome, Italy Search for more papers by this author Stefano M Cannata Corresponding Author Stefano M Cannata Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Dror Seliktar Dror Seliktar Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel Search for more papers by this author Giulio Cossu Corresponding Author Giulio Cossu Department of Cell and Developmental Biology, University College London, London, UK Institute of Inflammation and Repair, University of Manchester, Manchester, UK Search for more papers by this author Cesare Gargioli Corresponding Author Cesare Gargioli Department of Biology, Tor Vergata Rome University, Rome, Italy IRCCS MultiMedica, Milan, Italy Search for more papers by this author Claudia Fuoco Claudia Fuoco Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Roberto Rizzi Roberto Rizzi IRCCS MultiMedica, Milan, Italy Cell Biology and Neurobiology Institute, National Research Council of Italy, Rome, Italy Search for more papers by this author Antonella Biondo Antonella Biondo Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Emanuela Longa Emanuela Longa Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy Search for more papers by this author Anna Mascaro Anna Mascaro Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy Search for more papers by this author Keren Shapira-Schweitzer Keren Shapira-Schweitzer Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel Search for more papers by this author Olga Kossovar Olga Kossovar Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel Search for more papers by this author Sara Benedetti Sara Benedetti Department of Cell and Developmental Biology, University College London, London, UKCorrection added on 10 March 2015, after first online publication: author affiliations have been corrected. Search for more papers by this author Maria L Salvatori Maria L Salvatori Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Sabrina Santoleri Sabrina Santoleri Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Stefano Testa Stefano Testa Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Sergio Bernardini Sergio Bernardini Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Roberto Bottinelli Roberto Bottinelli Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy Fondazione Salvatore Maugeri (IRCCS), Scientific Institute of Pavia, Pavia, Italy Search for more papers by this author Claudia Bearzi Claudia Bearzi IRCCS MultiMedica, Milan, Italy Cell Biology and Neurobiology Institute, National Research Council of Italy, Rome, Italy Search for more papers by this author Stefano M Cannata Corresponding Author Stefano M Cannata Department of Biology, Tor Vergata Rome University, Rome, Italy Search for more papers by this author Dror Seliktar Dror Seliktar Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel Search for more papers by this author Giulio Cossu Corresponding Author Giulio Cossu Department of Cell and Developmental Biology, University College London, London, UK Institute of Inflammation and Repair, University of Manchester, Manchester, UK Search for more papers by this author Cesare Gargioli Corresponding Author Cesare Gargioli Department of Biology, Tor Vergata Rome University, Rome, Italy IRCCS MultiMedica, Milan, Italy Search for more papers by this author Author Information Claudia Fuoco1, Roberto Rizzi2,3, Antonella Biondo1, Emanuela Longa4, Anna Mascaro4, Keren Shapira-Schweitzer6, Olga Kossovar6, Sara Benedetti5, Maria L Salvatori1, Sabrina Santoleri1,9, Stefano Testa1, Sergio Bernardini1, Roberto Bottinelli4,7, Claudia Bearzi2,3, Stefano M Cannata 1, Dror Seliktar6, Giulio Cossu 5,8 and Cesare Gargioli 1,2 1Department of Biology, Tor Vergata Rome University, Rome, Italy 2IRCCS MultiMedica, Milan, Italy 3Cell Biology and Neurobiology Institute, National Research Council of Italy, Rome, Italy 4Department of Molecular Medicine and Interdepartmental Centre for Research in Sport Biology and Medicine, University of Pavia, Pavia, Italy 5Department of Cell and Developmental Biology, University College London, London, UK 6Faculty of Biomedical Engineering, Technion—Israel Institute of Technology, Haifa, Israel 7Fondazione Salvatore Maugeri (IRCCS), Scientific Institute of Pavia, Pavia, Italy 8Institute of Inflammation and Repair, University of Manchester, Manchester, UK 9Present address: Biocenter Oulu, Institute of Biomedicine, University of Oulu Finland *Corresponding author. Tel: +39 6 72594815; E-mail: [email protected] *Corresponding author. Tel: +44 161 3062526; E-mail: [email protected] *Corresponding author. Tel: +39 6 72594815; E-mail: [email protected] EMBO Mol Med (2015)7:411-422https://doi.org/10.15252/emmm.201404062 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Extensive loss of skeletal muscle tissue results in mutilations and severe loss of function. In vitro-generated artificial muscles undergo necrosis when transplanted in vivo before host angiogenesis may provide oxygen for fibre survival. Here, we report a novel strategy based upon the use of mouse or human mesoangioblasts encapsulated inside PEG-fibrinogen hydrogel. Once engineered to express placental-derived growth factor, mesoangioblasts attract host vessels and nerves, contributing to in vivo survival and maturation of newly formed myofibres. When the graft was implanted underneath the skin on the surface of the tibialis anterior, mature and aligned myofibres formed within several weeks as a complete and functional extra muscle. Moreover, replacing the ablated tibialis anterior with PEG-fibrinogen-embedded mesoangioblasts also resulted in an artificial muscle very similar to a normal tibialis anterior. This strategy opens the possibility for patient-specific muscle creation for a large number of pathological conditions involving muscle tissue wasting. Synopsis This study proposes a novel and efficient strategy for skeletal muscle tissue engineering based on in vitro mixing of mouse or human mesoangioblasts with a PEG-fibrinogen, which promotes their survival and differentiation into muscle. Upon graft implantation between the skin and the outer surface of the tibialis anterior, mature and aligned myofibres formed within a few weeks as a complete and functional extra muscle. Upon graft replacement of an almost completely ablated tibialis anterior, a new skeletal muscle very similar to the ablated one formed within a few weeks. Placental-derived growth factor-lentivirus transduction of muscle cells prior to transplant stimulates blood vessel growth and contributes to in vivo cells survival and maturation. This novel strategy opens up the possibility for patient-specific muscle engineering in a large number of pathological conditions involving muscle tissue wasting. Introduction Tissue engineering aims to create a microenvironment similar to the one where organogenesis took place. This requires the combined use of progenitor cells and biomaterials that allow optimal cell-matrix interactions, thus promoting better engraftment, differentiation and, consequently, better tissue generation. Skin, bone and cartilage, and recently trachea and urinary bladder, are successful examples of this strategy (Horch et al, 2000; Koop et al, 2004; Vangsness et al, 2004; Bader & Macchiarini, 2010; Watanabe et al, 2011). In contrast, tissue engineering for skeletal muscle still represents a difficult task despite its potential for the treatment of a variety of pathological conditions including post-traumatic muscle damages, post-surgery tissue ablation and incontinent sphincters (Guettier-Sigrist et al, 1998; Rizzi et al, 2012). Three-dimensional skeletal muscle tissue has been developed exploiting the influence of mechanical stretch (Vandenburgh & Kaufman, 1979), revealing the pivotal role played by kinetic forces on myofibre organization and muscle maturation. For example, Powell and colleagues (Powell et al, 2002) improved engineered three-dimensional (3D) human skeletal muscle on a collagen and matrigel scaffold by applying mechanical stimulation, thus increasing elasticity, and cross-sectional area (CSA). Moreover, several other recent developments, using decellularized natural scaffold to repair massive muscle injury, exploit host stem cells regenerative capabilities. The latter approach has shown only partial integration with damaged muscle and limited vascularization and innervation at the interface between the artificial and the host muscle (Corona et al, 2012; Sicari et al, 2012). Hence, this method still needs optimization, especially in terms of supporting blood vessel and nerves for artificial tissue survival and function. To overcome this hurdle, we looked for an innovative strategy exploiting a biomaterial able to promote myogenic cell differentiation in vivo so that angiogenesis and innervation may occur during muscle fibre formation and maturation. Results PEG-fibrinogen enhances Mabs myogenic differentiation A photopolymerizable hydrogel based upon polyethyleneglycol (PEG) and fibrinogen (PEG-fibrinogen: PF) (Almany & Seliktar, 2005; Fuoco et al, 2012; Seliktar, 2012) was combined with vessel-associated muscle progenitors, termed mesoangioblasts (Mabs), which are able to undergo robust myogenesis in vivo and in vitro. PF, which is clinically approved, creates a favourable microenvironment by coupling natural and synthetic features, adjustable to the needs of each specific cell type; Mabs are also already clinically approved, and a clinical trial (Eudract Number: 2011-000176-33) for Duchenne Muscular Dystrophy has been recently completed (G. Cossu et al, in preparation). The adult mouse C57 Mabs (Díaz-Manera et al, 2010) embedded into PF showed a remarkable muscle differentiation (Fig 1A and B) and were therefore selected for subsequent experiments and indicated as mouse Mabs (mMabs). A cylindrical silicon mould 0.5 cm high × 0.2 cm diameter was loaded with 100 μl solution of 8 mg/ml PF with 5 × 105 mMabs transduced with a lentiviral vector encoding the reporter gene nuclear β-galactosidase (mMabs-nLacZ); the constructs were formed by exposing the moulds to non-toxic long-wave UV light, as described in the Materials and Methods. The constructs were cultured for 24 h in serum-supplemented growth medium and then transferred into serum-depleted differentiation medium for 5 days in order to promote muscle fibre formation (Fig 1A and B). The Mabs rapidly underwent cell fusion in the PF within 24 h as documented by the time course performed in parallel with mMabs cultured into 2D standard plastic culture (TCP) versus mMabs encapsulated into PF (Supplementary Fig S1). Mabs moved well inside the gel (Supplementary Movie S1) and showed a much faster differentiation in the 3D PF cultures (Supplementary Fig S1D–F) than TCP one (Supplementary Fig S1A–C), leading to the formation of mature well-differentiated contractile myotubes. Of notice, even newly formed myotubes contracted spontaneously (Supplementary Movie S2) something usually not observed on TCP. We did not quantify proliferation and differentiation inside PF, as compared to control cells grown on plastic, due to the difficulty counting cells in the multiple focal plans within the gel. However, after 5 days in vitro, immunofluorescence analysis for myosin heavy chain (MyHC) and Western blot analysis revealed robust expression of skeletal muscle-specific proteins such as tropomyosin, troponin, creatine kinase and MyHC (Fig 1C and D). Furthermore, after 15 days of 3D culture into 8 mg/ml PF, mMabs generated a very thick three-dimensional network of fibres able to contract the plug as whole (Supplementary Movie S3). Figure 1. In vitro characterization of Mabs grown inside PF hydrogels A. Phase contrast images of Mabs cultured into PF hydrogels (5 days): cells formed a thick three-dimensional myotube network with numerous thick myofibres (arrows). Scale bar: 50 μm. B. Confocal immunofluorescence analysis with an antibody against myosin heavy chain (MyHC) (red) and SYTOX green which labels nuclei (green), showing multi-nucleated, mature muscle fibres (arrows) developed from Mabs in PF (5 days). Scale bar: 50 μm. C, D. Western blot and densitometry analysis of different protein extracts of Mabs differentiated inside PF constructs (PF) showing expression levels comparable to adult TA (Ctrl). Group of n = 6 of Mabs differentiated inside PF and host TA has been tested, densitometric analysis from n = 3 different Western blots gauging the level of expression of muscle markers is presented as means ± standard error. Source data are available online for this figure. Source Data for Figure 1 [emmm201404062-sup-0007-source_data_Fig1.pdf] Download figure Download PowerPoint PlGF promotes in vivo vessel recruitment Undifferentiated mMabs embedded in PF were subsequently implanted under the back skin of 2-month-old RAG2/γchain null mice (Cao et al, 1995). Immunodeficient mice were used to prevent immune reaction against the bacterial LacZ gene used as reporter. Angiogenesis in the implanted tissue was enhanced by transduction of the mMabs-nLacZ with a lentiviral vector expressing placental-derived growth factor (PlGF) that we had previously shown to enhance angiogenesis of transplanted mMabs (Gargioli et al, 2008). To rule out the possibility that PlGF expression may interfere with mMabs differentiation, we performed immunofluorescence analysis for MyHC which showed similar myogenic differentiation of both the wild-type mMabs and mMabs-nLacZ expressing PlGF (mMabs-nLacZ/PlGF) (Supplementary Fig S2A and B). In vivo analysis of dorsal subcutaneous PF implants loaded with 1.5 × 106 mMabs-nLacZ/PlGF revealed increased blood vessel density in comparison with control mMabs-nLacZ (Supplementary Fig S2C–G), quantified by counting the number of blood vessels, stained for VE-cadherin per muscle fibre (Supplementary Fig S2H). Robust muscle differentiation of cells embedded into PF was observed, with enhanced differentiation in mMabs-nLacZ/PlGF comparing with unmodified mMabs (Supplementary Fig S2I–L). The mMabs-nLacZ/PlGF showed an increased number of MyHC-positive myofibres in the centre of the implant due to an enhanced vascularization and thus to improved oxygenation and nutrition, while unmodified mMabs generated fewer and smaller muscle fibres in the centre of the implant (Supplementary Fig S2I and J). Because of these results, only mMabs expressing PlGF were utilized for subsequent in vivo studies. Hydrogel resorption was analysed in subcutaneous implants of 1.5 × 106 mMabs-nLacZ/PlGF encapsulated into 50 μl of 8 mg/ml PF: after 3 days, the PF regular structure still surrounded the cells, whereas after 7 days, the PF was almost completely resorbed (Supplementary Fig S2M and N). In both conditions (with or without PlGF), newly formed myofibre was randomly oriented and this did not result in any structure anatomically recognizable as a skeletal muscle (Supplementary Fig S2K and L). Contracting muscle surface as 'anatomical bioreactor' promoting fibres alignment We reasoned that a functional, contractile muscle could influence the implant evoking myofibres alignment in response to its contractile activity mimicking a stretching stimulus. Thus, we implanted mMabs-nLacZ/PlGF (1.5 × 106) in PF constructs (cylindrically shaped, 0.5 cm high × 0.2 cm diameter) under the skin covering the surface of the tibialis anterior (TA) to exploit its contractile activity and then promoting artificial muscle fibres alignment recapitulating the normal skeletal muscle tissue architecture (Supplementary Fig S3 and Fig 2A). The resulting structures were collected at early (4 weeks) and late stages (8 weeks) after implantation. The implants were completely incorporated by the host TA epimysium (Fig 2B) and were revealed only when tendons were cut (Fig 2C). The resulting structure showed size and morphology very similar to the underlying TA muscle (Fig 2C and D). Macroscopic observation of the whole muscle showed a network of blood vessels on the muscle-like tissue (Fig 2E), while systemic ink injection via recipient mouse femoral artery confirmed the expected connection with the host vascular tree (Fig 2F). Western blot analysis of muscle and vessel protein expression was performed on crude extracts from 8-week implanted tissue and revealed an expression pattern comparable to control host TA (Fig 2G and H). The development of a mature and vascularized artificial muscle tissue was further confirmed by real-time quantitative PCR, which revealed comparable expression levels of all the muscle-specific genes examined (Supplementary Fig S4). β-Galactosidase staining on histological sections of early (4 weeks) and mature (8 weeks) artificial muscle revealed LacZ-positive donor nuclei centrally located at early stages but peripherally located at 8 weeks, whereas cells derived from the host were located mainly in the interstitial structures where the LacZ-positive cells were mostly absent (Fig 2I–L). Immunofluorescence analysis with anti-laminin antibodies revealed an incomplete basal lamina and residual interstitial tissue at 4 weeks; in contrast, at 8 weeks, the artificial muscle showed an almost normal muscle tissue organization with well-patterned basal lamina (Fig 2M–P). Immunostaining for neurofilaments and bungarotoxin staining (which binds the acetylcholine receptor) (Supplementary Fig S5A) were performed at early and late stages and showed a progressive maturation of the developing synapses; only axons were detected at early stages (Supplementary Fig S5B), whereas bungarotoxin-positive, neuromuscular plaques were detected at late stages (Supplementary Fig S5C). Neuromuscular synapses were also labelled by immunofluorescence and visualized by confocal microscopy; the isosurfaces showed the formation of mature neuromuscular plaques in the generated artificial new muscle (Fig 2Q). Moreover, the artificial muscle contained many small vessels identified by immunostaining with antibodies against smooth muscle actin (SMA) and vascular endothelial cadherin (VE-Cad) (Fig 2R). Despite an apparently normal muscular morphology and gene expression pattern (Supplementary Fig S4), careful examination of the artificial muscle and histological longitudinal sections did not reveal tendons at the extremities (Fig 2S). Interestingly, several nLacZ donor nuclei were detected in the underlying TA—which was not injured during the implantation—confirming that myogenic cells may colonize adjacent muscles in small rodents (Hughes & Blau, 1990). This finding may also explain why only a part of the artificial muscle had LacZ-positive nuclei, due to incomplete transduction (about 80%) of donor mesoangioblasts and possible cell migration from the underlying TA (Supplementary Fig S6A–F). Moreover, the new muscle tissue showed re-constituted Pax7-positive satellite cell pool, in part expressing β-galactosidase (Supplementary Fig S6G–I). Therefore, in order to verify host contribution to artificial tissue generation, we performed experiments into ubiquitous GFP mice background using PF-embedded mMabs-PlGF. Because of an immune response against implanted PF and mMabs-derived structure, we could analyse only short-time engraftment (15 days), when we observed myogenic differentiation of implanted mMabs in the absence of GFP-positive cells that could be detected in the adjacent tissue (Supplementary Fig S7). Furthermore, the artificial muscle ability to regenerate was investigated by cardiotoxin injection that caused myofibre degeneration, followed by muscle fibre regeneration with kinetics comparable to that of a normal TA (Supplementary Fig S8). After damage, the artificial muscle and the underlying TA contain satellite cells with similar myogenic potency: magnetically sorted satellite cells showed LacZ expression when isolated from the artificial muscle but not from the host TA and underwent myogenesis with comparable efficiency (Supplementary Fig S9 and Supplementary Table S1). Finally, in order to understand whether artificial muscle would respond to hypertrophic or atrophic stimuli like a normal muscle, clenbuterol administration and denervation were performed (Supplementary Fig S10). The analysis revealed a striking atrophic response to sciatic nerve resection both in the artificial muscle and in the underlying TA (Supplementary Fig S10G–J), while the clenbuterol treatment showed only a modest, which did not reach statistical significance in both artificial and underlying muscles, possibly due to inadequate response time or concentration (Supplementary Fig S10C–F); quantification obtained evaluating CSA average of the treated artificial and TA muscles in ten non-adjacent sections (Supplementary Fig S10A and B). Figure 2. Implanted constructs of Mabs in PF on the surface of TA A. Gross morphology of the PF implant containing Mabs-nLacZ/PlGF immediately after implantation on the surface of underlying TA (arrow). B, C. Artificial muscles (arrows) developed within 8 weeks over the host TA surface (arrowheads), encapsulated by the host perimysium (B) and released after tendon resection of the TA (C). D. The isolated artificial muscle from the TA surface (arrow) is comparable in size to the underlying host TA (arrowhead). E. Stereomicroscopy images revealing blood vessel formation in the artificial muscle. Scale bar: 100 μm. F. X-Gal and H&E staining on artificial muscle section revealing vessel connection with host vasculature by Indian ink, injected into the femoral artery immediately before sacrifice. Scale bar: 20 μm. G. Western blot analysis of protein extracts from artificial muscles (PF) and host TA (Ctrl) reveals remarkable expression of muscle- and vessel-specific proteins (n = 5). H. Densitometric analysis from n = 3 different Western blots gauging the level of expression of muscle- and vessel-specific proteins; presented values are expressed as means ± standard error. I–L. X-Gal and H&E staining of artificial muscle sections at 4 (I, K) and 8 weeks (J, L) after implantation, showing donor origin of muscle cells (nLacZ+ nuclei) and correlation between implant duration and myofibre maturation (I, J images are the results of the collage of several photos). Scale bar: (I, J) 200 μm, (K, L) 100 μm. M–P. Immunofluorescence analysis on adjacent sections revealing laminin organization (green) and myosin heavy chain (MyHC) expression (red) at 4 weeks (M) and 8 weeks (N) after grafting (M, N displayed items are obtained from a multi-photo collage); DAPI staining (blue) shows centrally located nuclei in developing fibres (O) and peripheral nuclei in mature myofibres (P). Scale bar: (M, N) 200 μm, (O, P) 50 μm. Q. Confocal isosurface image of neurofilament (green) and bungarotoxin (red) immunofluorescence in the artificial muscle section shows a mature neuromuscular plaque. Scale bar: 10 μm. R. Immunofluorescence against smooth muscle actin (blue), laminin (green) and VE-Cad (red) superimposed on phase contrast image shows blood vessels (arrows) adjacent to the myofibres in the artificial muscle section. Scale bar: 20 μm. S. X-Gal staining on mature artificial muscle (arrowhead) and the underneath host TA (arrow) revealing no tendon development at the extremity of the artificial tissue; implanted LacZ-positive Mabs are present within the host TA. Scale bar: 200 μm. Source data are available online for this figure. Source Data for Figure 2 [emmm201404062-sup-0008-source_data_Fig2.pdf] Download figure Download PowerPoint Artificial muscle closely resembles a natural skeletal muscle Ultrastructural and functional analyses were conducted to evaluate to what extent the artificial muscle resembles a normal adult muscle. Electron microscopy analysis revealed a completed sarcomerogenesis typical of mature skeletal muscle while staining with blu-O-gal revealed the presence of donor-derived LacZ-positive nuclei adjacent to transversely sectioned sarcomeres (Fig 3A and B). Single artificial myofibres were isolated from bunches of artificial muscle fibres (LacZ-positive) and showed normal cross-striation (Fig 3C and D). A subset of the artificial muscle fibres (n = 20) was stained by immunofluorescence with antibodies against β-galactosidase to assess the origin of their nuclei: results showed that the large majority of nuclei were of donor origin (based on nuclear LacZ expression), but few unlabelled nuclei, whose origin remain unclear, were also present (Fig 3E and F) (Hughes & Blau, 1990). A large number of nLacZ-positive artificial muscle fibres (n = 153) and underlying TA muscle control fibres (n = 103) from 4-month-old C57 RAG2/γchain−/− mice were mounted in a set-up which enabled viewing their striation pattern at 320-times magnification, determining CSA, specific force (Po/CSA) and maximum shortening velocity (Vo) (Bottinelli et al, 1996; Pellegrino et al, 2003). The striation pattern, CSA, Po/CSA and Vo of muscle fibres from artificial muscle and from control muscle were indistinguishable (Fig 3G–I). Po/CSA values were similar in single fibres from the artificial muscle and the underlying control muscle. Moreover, the values of specific force reported in Fig 3G are fully consistent with our previous findings on muscle fibres from normal TA muscles of mice (Torrente et al, 2004). Likewise, the value of Vo, which is mainly dependent on the MyHC isoform content, indicated that the kinetic of actomyosin interaction was virtually identical to that of the normal adult fibres of the underlying TA (Bottinelli et al, 1994; Pellegrino et al, 2003; Torrente et al, 2004). At the end of the functional analysis, the MyHC isoform content of each fibre was also determined by SDS–PAGE. All fibres analysed were found to contain adult MyHC isoforms and almost exclusively MyHC-2B, which is the most expressed MyHC isoform in fast mouse muscles (Pellegrino et al, 2003) (Fig 3J and K). Figure 3. Structural and functional analysis of artificial muscle and underlying TA control muscle A. Electron microscopy with Bluo-Gal staining reveals a LacZ-positive donor nucleus (arrow) peripheral to trans