Reversible immortalisation enables genetic correction of human muscle progenitors and engineering of next‐generation human artificial chromosomes for Duchenne muscular dystrophy
2017; Springer Nature; Volume: 10; Issue: 2 Linguagem: Inglês
10.15252/emmm.201607284
ISSN1757-4684
AutoresSara Benedetti, Narumi Uno, Hidetoshi Hoshiya, Martina Ragazzi, Giuliana Ferrari, Yasuhiro Kazuki, Louise A. Moyle, Rossana Tonlorenzi, Angelo Lombardo, Soraya Chaouch, Vincent Mouly, Marc T. Moore, Linda Popplewell, Kanako Kazuki, Motonobu Katoh, Luigi Naldini, George Dickson, Graziella Messina, Mitsuo Oshimura, Giulio Cossu, Francesco Saverio Tedesco,
Tópico(s)Pluripotent Stem Cells Research
ResumoResearch Article14 December 2017Open Access Source DataTransparent process Reversible immortalisation enables genetic correction of human muscle progenitors and engineering of next-generation human artificial chromosomes for Duchenne muscular dystrophy Sara Benedetti Sara Benedetti Department of Cell and Developmental Biology, University College London, London, UK Great Ormond Street Institute of Child Health, University College London, London, UK Search for more papers by this author Narumi Uno Narumi Uno Department of Biomedical Science, Institute of Regenerative Medicine and Biofunction, Tottori University, Yonago, Tottori, Japan Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Hidetoshi Hoshiya Hidetoshi Hoshiya Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Martina Ragazzi Martina Ragazzi Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Giulia Ferrari Giulia Ferrari Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Yasuhiro Kazuki Yasuhiro Kazuki Department of Biomedical Science, Institute of Regenerative Medicine and Biofunction, Tottori University, Yonago, Tottori, Japan Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Louise Anne Moyle Louise Anne Moyle Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Rossana Tonlorenzi Rossana Tonlorenzi Division of Neuroscience, Institute of Experimental Neurology, San Raffaele Scientific Institute, Milan, Italy Search for more papers by this author Angelo Lombardo Angelo Lombardo San Raffaele Telethon Institute for Gene Therapy (TIGET), San Raffaele Scientific Institute and Vita Salute San Raffaele University, Milan, Italy Search for more papers by this author Soraya Chaouch Soraya Chaouch AIM/AFM Center for Research in Myology, Sorbonne Universités, UPMC Univ. Paris 06, INSERM UMRS974, CNRS FRE3617, Paris, France Search for more papers by this author Vincent Mouly Vincent Mouly AIM/AFM Center for Research in Myology, Sorbonne Universités, UPMC Univ. Paris 06, INSERM UMRS974, CNRS FRE3617, Paris, France Search for more papers by this author Marc Moore Marc Moore School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK Search for more papers by this author Linda Popplewell Linda Popplewell School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK Search for more papers by this author Kanako Kazuki Kanako Kazuki Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Motonobu Katoh Motonobu Katoh Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Luigi Naldini Luigi Naldini orcid.org/0000-0002-7835-527X Department of Biosciences, University of Milan, Milan, Italy Search for more papers by this author George Dickson George Dickson School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK Search for more papers by this author Graziella Messina Graziella Messina Department of Biosciences, University of Milan, Milan, Italy Search for more papers by this author Mitsuo Oshimura Mitsuo Oshimura Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Giulio Cossu Corresponding Author Giulio Cossu [email protected] orcid.org/0000-0001-5863-9593 Division of Cell Matrix Biology and Regenerative Medicine, University of Manchester, Manchester, UK Search for more papers by this author Francesco Saverio Tedesco Corresponding Author Francesco Saverio Tedesco [email protected] orcid.org/0000-0001-5321-7682 Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Sara Benedetti Sara Benedetti Department of Cell and Developmental Biology, University College London, London, UK Great Ormond Street Institute of Child Health, University College London, London, UK Search for more papers by this author Narumi Uno Narumi Uno Department of Biomedical Science, Institute of Regenerative Medicine and Biofunction, Tottori University, Yonago, Tottori, Japan Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Hidetoshi Hoshiya Hidetoshi Hoshiya Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Martina Ragazzi Martina Ragazzi Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Giulia Ferrari Giulia Ferrari Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Yasuhiro Kazuki Yasuhiro Kazuki Department of Biomedical Science, Institute of Regenerative Medicine and Biofunction, Tottori University, Yonago, Tottori, Japan Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Louise Anne Moyle Louise Anne Moyle Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Rossana Tonlorenzi Rossana Tonlorenzi Division of Neuroscience, Institute of Experimental Neurology, San Raffaele Scientific Institute, Milan, Italy Search for more papers by this author Angelo Lombardo Angelo Lombardo San Raffaele Telethon Institute for Gene Therapy (TIGET), San Raffaele Scientific Institute and Vita Salute San Raffaele University, Milan, Italy Search for more papers by this author Soraya Chaouch Soraya Chaouch AIM/AFM Center for Research in Myology, Sorbonne Universités, UPMC Univ. Paris 06, INSERM UMRS974, CNRS FRE3617, Paris, France Search for more papers by this author Vincent Mouly Vincent Mouly AIM/AFM Center for Research in Myology, Sorbonne Universités, UPMC Univ. Paris 06, INSERM UMRS974, CNRS FRE3617, Paris, France Search for more papers by this author Marc Moore Marc Moore School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK Search for more papers by this author Linda Popplewell Linda Popplewell School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK Search for more papers by this author Kanako Kazuki Kanako Kazuki Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Motonobu Katoh Motonobu Katoh Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Luigi Naldini Luigi Naldini orcid.org/0000-0002-7835-527X Department of Biosciences, University of Milan, Milan, Italy Search for more papers by this author George Dickson George Dickson School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK Search for more papers by this author Graziella Messina Graziella Messina Department of Biosciences, University of Milan, Milan, Italy Search for more papers by this author Mitsuo Oshimura Mitsuo Oshimura Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan Search for more papers by this author Giulio Cossu Corresponding Author Giulio Cossu [email protected] orcid.org/0000-0001-5863-9593 Division of Cell Matrix Biology and Regenerative Medicine, University of Manchester, Manchester, UK Search for more papers by this author Francesco Saverio Tedesco Corresponding Author Francesco Saverio Tedesco [email protected] orcid.org/0000-0001-5321-7682 Department of Cell and Developmental Biology, University College London, London, UK Search for more papers by this author Author Information Sara Benedetti1,2, Narumi Uno3,4,‡, Hidetoshi Hoshiya1,11,‡, Martina Ragazzi1,12, Giulia Ferrari1, Yasuhiro Kazuki3,4, Louise Anne Moyle1, Rossana Tonlorenzi5, Angelo Lombardo6, Soraya Chaouch7, Vincent Mouly7, Marc Moore8, Linda Popplewell8, Kanako Kazuki4, Motonobu Katoh4, Luigi Naldini9, George Dickson8, Graziella Messina9,‡, Mitsuo Oshimura4,‡, Giulio Cossu *,10,‡ and Francesco Saverio Tedesco *,1,‡ 1Department of Cell and Developmental Biology, University College London, London, UK 2Great Ormond Street Institute of Child Health, University College London, London, UK 3Department of Biomedical Science, Institute of Regenerative Medicine and Biofunction, Tottori University, Yonago, Tottori, Japan 4Chromosome Engineering Research Center (CERC), Tottori University, Yonago, Tottori, Japan 5Division of Neuroscience, Institute of Experimental Neurology, San Raffaele Scientific Institute, Milan, Italy 6San Raffaele Telethon Institute for Gene Therapy (TIGET), San Raffaele Scientific Institute and Vita Salute San Raffaele University, Milan, Italy 7AIM/AFM Center for Research in Myology, Sorbonne Universités, UPMC Univ. Paris 06, INSERM UMRS974, CNRS FRE3617, Paris, France 8School of Biological Sciences, Royal Holloway-University of London, Egham, Surrey, UK 9Department of Biosciences, University of Milan, Milan, Italy 10Division of Cell Matrix Biology and Regenerative Medicine, University of Manchester, Manchester, UK 11Present address: Cell and Gene Therapy Catapult, Guy's Hospital, Great Maze Pound, London, UK 12Present address: MolMed S.p.A, Milan, Italy ‡These authors contributed equally to this work ‡These authors contributed equally to this work as senior authors *Corresponding author. Tel: +44 161 3062526; E-mail: [email protected] *Corresponding author. Tel: +44 2031 082383; E-mail: [email protected] EMBO Mol Med (2018)10:254-275https://doi.org/10.15252/emmm.201607284 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 Transferring large or multiple genes into primary human stem/progenitor cells is challenged by restrictions in vector capacity, and this hurdle limits the success of gene therapy. A paradigm is Duchenne muscular dystrophy (DMD), an incurable disorder caused by mutations in the largest human gene: dystrophin. The combination of large-capacity vectors, such as human artificial chromosomes (HACs), with stem/progenitor cells may overcome this limitation. We previously reported amelioration of the dystrophic phenotype in mice transplanted with murine muscle progenitors containing a HAC with the entire dystrophin locus (DYS-HAC). However, translation of this strategy to human muscle progenitors requires extension of their proliferative potential to withstand clonal cell expansion after HAC transfer. Here, we show that reversible cell immortalisation mediated by lentivirally delivered excisable hTERT and Bmi1 transgenes extended cell proliferation, enabling transfer of a novel DYS-HAC into DMD satellite cell-derived myoblasts and perivascular cell-derived mesoangioblasts. Genetically corrected cells maintained a stable karyotype, did not undergo tumorigenic transformation and retained their migration ability. Cells remained myogenic in vitro (spontaneously or upon MyoD induction) and engrafted murine skeletal muscle upon transplantation. Finally, we combined the aforementioned functions into a next-generation HAC capable of delivering reversible immortalisation, complete genetic correction, additional dystrophin expression, inducible differentiation and controllable cell death. This work establishes a novel platform for complex gene transfer into clinically relevant human muscle progenitors for DMD gene therapy. Synopsis Reversible immortalisation of human dystrophic muscle progenitors with a novel human artificial chromosome (HAC) containing the entire dystrophin locus (DYS-HAC2) enables genetic correction & provide evidence of translation of HAC technology into DMD muscle progenitors for ex vivo gene therapy. Lentivirally delivered hTERT and Bmi1 cDNAs extend proliferation of human muscle progenitors (both myoblasts and pericyte-derived mesoangioblasts) and can be reverted by administering Cre recombinase and ganciclovir. Extension of the proliferative ability of muscle progenitors derived from Duchenne muscular dystrophy patients enables their genetic correction with a novel DYS-HAC. A next-generation DYS-HAC containing multiple gene functions was engineered to enable simultaneous delivery of: (i) genomic integration-free reversible immortalisation, (ii) genetic correction, (iii) inducible differentiation, (iv) controllable cell death. Introduction Duchenne muscular dystrophy (DMD) is the most common muscle disorder of childhood and one of the most severe forms of muscular dystrophy that leads to progressive muscle wasting and premature death (Mercuri & Muntoni, 2013a). DMD is caused by mutations in the X-linked dystrophin gene, which encodes for a protein responsible for sarcolemma integrity (Hoffman et al, 1987). Despite extensive pre-clinical work and many novel clinical trials (Benedetti et al, 2013; Mercuri & Muntoni, 2013b; Bengtsson et al, 2016), currently there are no definitive treatments. One of the main obstacles to the development of an effective gene therapy for DMD is the large size of the dystrophin gene (2.4 Mb), preventing cloning of its full cDNA (14 kb) into conventional gene therapy vectors. This limitation is then amplified by the abundance of degenerating skeletal muscle needing repair or regeneration. In recent years, a new generation of large cloning capacity gene delivery vectors named human artificial chromosomes (HACs) has been developed (Kazuki & Oshimura, 2011; Kouprina et al, 2014; Oshimura et al, 2015; Tedesco, 2015). HACs present several advantages for DMD gene therapy, including (i) the ability to carry large DNA sequences such as entire genetic loci, thus resulting in a more physiological expression of genes with complex transcriptional regulation (including dystrophin), (ii) stable episomal maintenance of a single copy gene, avoiding the risk of insertional oncogenesis. In this context, a HAC containing the entire dystrophin locus (DYS-HAC) was engineered for ex vivo stem cell gene therapy of DMD (Hoshiya et al, 2009). In the last two decades, several populations of stem/progenitor cells with myogenic potential have been isolated (Tedesco et al, 2010, 2017), but so far only satellite cell-derived myoblasts (Perie et al, 2014; Skuk & Tremblay, 2014), muscle pericyte-derived mesoangioblasts (Cossu et al, 2015) and (to a minor extent) muscle-derived AC133+ cells (Torrente et al, 2007) have undergone clinical experimentation. We have shown that transplantation of murine dystrophic mesoangioblasts corrected with a DYS-HAC ameliorated the phenotype of dystrophic mdx mice (Tedesco et al, 2011). These results provided the first evidence of safe and efficacious pre-clinical gene replacement therapy with a HAC into an animal model of a genetic disease, paving the way for translating HAC gene transfer to human cells. However, proliferation of human and murine cells is regulated by different pathways, often resulting in a limited lifespan of human somatic cells. Therefore, primary human muscle cells are likely to require an extension of their proliferative potential to withstand selection of corrected cells and subsequent expansion to clinically relevant numbers after clonal HAC transfer [in the range of 109 cells (Cossu et al, 2015)]. Here, we developed a novel genetic correction strategy based upon the use of reversibly immortalising lentiviral vectors to extend the proliferative potential of human myoblasts and mesoangioblasts. We show that DMD muscle progenitor cells immortalised by means of lentiviral vectors expressing the excisable catalytic subunit of human telomerase (hTERT) and the cell cycle regulator Bmi1 (Cudre-Mauroux et al, 2003) enable transfer of a novel DYS-HAC. Moreover, the presence of the herpes simplex virus thymidine kinase (HSV-TK) cDNA between loxP sites allows selective drug-induced elimination of target cells that escaped transgene excision by Cre recombination (Salmon et al, 2000). DMD, reversibly immortalised, DYS-HAC-corrected progenitors were transplanted in mouse models of acute and chronic muscle injury, where they engrafted regenerating skeletal muscle. Lastly, we combined all relevant gene functions into a single next-generation synthetic HAC capable of delivering reversible immortalisation, complete genetic correction, additional dystrophin expression, inducible differentiation and controllable cell death, generating the largest and possibly most complex gene therapy vector developed to date. Results Generation of a novel human artificial chromosome containing the entire human dystrophin locus To facilitate pre-clinical development of the DYS-HAC platform for human myogenic cells, we engineered a novel DYS-HAC devoid of potentially immunogenic gene products such as the enhanced green fluorescent protein (EGFP), blasticidin (Bsd), HSV1-TK (Tk) and hypoxanthine-guanine phosphoribosyltransferase (HPRT). For simplicity, the DYS-HAC previously engineered (Hoshiya et al, 2009) and used in our former murine study (Tedesco et al, 2011) is here renamed DYS-HAC1, to distinguish it from the newly generated DYS-HAC2. DYS-HAC2 was engineered by homologous recombination-mediated DYS-HAC1 modifications (Fig 1A). In order to remove EGFP, Bsd, HPRT and HSV1-TK genes from DYS-HAC1 and to add the floxable (FRT) neomycin (Neo) gene for selection and lox71 site and 5′ HPRT for further gene insertion, the targeting vector pN (Fig 1A), containing both the Neo gene and the regions for homologous recombination (A: 3.8 kb and B: 2.6 kb), was introduced by electroporation into chicken lymphoid DT40 cells already containing DYS-HAC1 [DT40(DYS-HAC1)]. DT40 hybrids were then subjected to G418 (neomycin) selection, and 19 G418-resistant DT40 hybrids were randomly selected for further analysis. One clone was found to be EGFP negative (Fig 1B), indicating successful homologous recombination. PCRs using primers able to discriminate DYS-HAC2 from DYS-HAC1 confirmed correct targeting (Fig 1C), whereas another set of primers designed on the dystrophin gene confirmed sequence integrity (Fig 1D). Finally, FISH analysis established that DYS-HAC2 segregated independently without host genome insertions or translocations (Fig 1E). Figure 1. Generation of a novel HAC containing the entire human dystrophin locus by homologous recombination The scheme shows a linearised map of the vectors and the strategy used to generate DYS-HAC2 by homologous recombination of DYS-HAC1 (Hoshiya et al, 2009). pN targeting vector, which contains regions for homologous recombination (A: 3.8 kb and B: 2.6 kb, in green) and a floxable (FRT) neomycin (Neo), was used to remove extra genes (EGFP, Bsd, HPRT and Tk) on DYS-HAC1 and to insert a floxable Neo gene. Primers designed to amplify DYS-HAC1 or DYS-HAC2 specific regions are highlighted in red. Phase contrast (left) and fluorescence (EGFP, right) images of DT40 cells containing DYS-HAC1 and DYS-HAC2. Scale bar: 50 μm. PCR analyses to discriminate between DYS-HAC1 and DYS-HAC2. DT40 cells: negative control. PCR panel to detect dystrophin exons in DT40(DYS-HAC1) and DT40(DYS-HAC2) cells. DT40 cells: negative control; human mesoangioblasts: positive control. In situ fluorescence hybridisation (FISH) analysis of DT40(DYS-HAC2) cells. White arrowheads: DYS-HAC2. Red: rhodamine-human COT-1 DNA; green: dystrophin FITC-DMD-BAC RP11-954B16; yellow: merge. Scale bar: 5 μm. DT40(DYS-HAC2) hybrid was used to transfer the DYS-HAC2 in CHO cells (complete list in Appendix Table S1). FISH analyses of CHO(DYS-HAC2)-7 (left) and A9(DYS-HAC2)-9 (right) clones. White arrowheads: DYS-HAC2. CHO(DYS-HAC2) hybrid was used to transfer DYS-HAC2 in A9 cells (complete list in Appendix Table S2). Red/purple: rhodamine-human COT-1 DNA; green: dystrophin FITC-DMD-BAC RP11-954B16; yellow: merge. Scale bar: 5 μm. Source data are available online for this figure. Source Data for Figure 1 [emmm201607284-sup-0004-SDataFig1.jpg] Download figure Download PowerPoint The novel DYS-HAC2 was then transferred via microcell-mediated chromosome transfer (MMCT) from DT40(DYS-HAC2) cells into Chinese hamster ovary (CHO) cells, to scale-up production of microcells for HAC transfer. Twenty CHO(DYS-HAC2) clones were randomly picked after G418 selection and both PCR and FISH confirmed the presence of a single episomal copy of DYS-HAC2 in four clones (Appendix Table S1). Among them, clone CHO(DYS-HAC2)-7 was then selected for transfer of DYS-HAC2 into murine A9 cells, as they have an even higher efficiency in generating microcells compared to CHO cells. Following G418 selection, 27 A9(DYS-HAC2) clones were randomly picked and presence of DYS-HAC2 was confirmed by PCR in five clones. FISH analysis showed episomal presence of DYS-HAC2 in three out of five A9(DYS-HAC2) clones (Appendix Table S2). Figure 1F shows fluorescent in situ hybridisation (FISH) images of CHO(DYS-HAC2)-7 and A9(DYS-HAC2)-9 clones utilised as DYS-HAC2 donors in subsequent experiments. Reversible immortalisation of DMD myoblasts enables DYS-HAC transfer and complete genetic correction Combined expression of hTERT and Bmi1 was shown to immortalise human myoblasts (Cudre-Mauroux et al, 2003). To test the hypothesis that reversible immortalisation could extend cell proliferation enough to allow HAC transfer in human myogenic progenitors, we planned to transfer the newly generated DYS-HAC2 (Fig 1) into reversibly immortalised DMD myoblasts (riDMD myoblasts). Before proceeding with HAC transfer, we confirmed that riDMD myoblasts (i) contained and transcribed hTERT and Bmi1 transgenes (Fig EV1A), (ii) had maintained their myogenic potential (Fig EV1B), (iii) generated dystrophin-deficient myotubes in vitro (Fig EV1C), (iv) were not tumorigenic (N = 3; Table EV1). We then transferred DYS-HAC2 into riDMD myoblasts via MMCT. Four G418-resistant clones, namely riDMD(DYS-HAC2)#α, #β, #γ and #δ, were selected. PCR analysis showed that two out of the four clones were positive for all analysed HAC regions (clones #α and #δ; Fig 2A, red boxes). Parental riDMD myoblasts have a deletion from exon 5 to exon 7 in the dystrophin gene (Fig 2B, first lane), which would result into an out-of-frame mutation: this was indeed confirmed by dystrophin transcript analysis, which demonstrated an out-of-frame mutation in riDMD myoblasts and ruled out a potential restoration of the reading frame by skipping of exon 8 (Fig EV1D; Muntoni et al, 1994; Cudre-Mauroux et al, 2003). Importantly, PCRs for dystrophin exons 5, 6 and 7 after HAC transfer demonstrated that both riDMD(DYS-HAC2)#α and #δ were positive for the exons originally deleted in the parental DMD myoblasts (Fig 2B), showing correction of the dystrophin gene sequence. FISH and karyotype analyses confirmed the presence of a single copy of DYS-HAC2 and a normal karyotype (Fig 2C). Subcutaneous injection of DYS-HAC2-corrected DMD myoblasts into immunodeficient scid/beige mice (N = 5 per clone) did not result in tumour formation (N = 10; Table EV1). Finally, riDMD(DYS-HAC2)#α myoblasts showed restoration of dystrophin mRNA expression and protein production in skeletal myotubes upon in vitro differentiation (Fig 2D–F; detailed analysis of myogenic differentiation in Appendix Fig S1A). Click here to expand this figure. Figure EV1. Characterisation of DMD immortalised (riDMD) myoblasts PCRs for hTERT and Bmi1 on genomic DNA and cDNA of reversibly immortalised myoblasts (riDMD myoblasts). Positive control: immortalised mesoangioblasts. riDMD myoblasts in proliferation (phase contrast, upper images) and after myogenic differentiation (lower images). Red: myosin heavy chain (MyHC); blue: Hoechst. Scale bar: 100 μm. Dystrophin immunofluorescence in riDMD myoblasts myotubes (white arrowheads). Red: MyHC; green: dystrophin; blue: Hoechst; yellow: merge. Scale bar: 50 μm. RT–PCR for dystrophin exon 3–9 transcript in differentiated riDMD myoblasts (deletion exons 5–7) confirming the presence of an out-of-frame DMD mutation and absence of alternative splicing variants (i.e. skipping of exon 8), which could potentially restore the reading frame. Healthy myoblasts: positive control. riDMD myoblast band is approximately 450 bp due to amplification of dystrophin exons 3, 4, 8 and 9, whereas healthy myoblast band is expected to be 833 bp due to amplification of exons 3, 4, 5, 6, 7, 8 and 9. Source data are available online for this figure. Download figure Download PowerPoint Figure 2. DYS-HAC2 transfer into reversibly immortalised DMD myoblasts PCR panel for HAC sequences in four neomycin-resistant riDMD(DYS-HAC2) myoblast clones. Red boxes: positive riDMD(DYS-HAC2) myoblasts. PCR analysis of human dystrophin exons on genomic DNA of riDMD myoblasts, riDMD(DYS-HAC2)#α and riDMD(DYS-HAC2)#δ clones. Parental riDMD myoblasts: negative control for exons 5–7; CHO(DYS-HAC2)-7: positive control. Left panel: karyotype analysis of riDMD myoblasts, riDMD(DYS-HAC2)#α and riDMD(DYS-HAC2)#δ myoblast clones. Right panel: FISH analysis on riDMD myoblasts and two selected riDMD(DYS-HAC2) myoblast clones (#α and #δ). Insets: magnifications showing single episomal DYS-HAC2. Red/purple: rhodamine-p11-4 human alpha satellite (centromeres of chromosome 13 and 21, hChr 13/21(cen)); green: dystrophin FITC-DMD-BAC RP11-954B16; yellow: merge. White arrowheads: DYS-HAC2. Scale bar: 3 μm. RT–PCR panel for dystrophin expression in differentiated riDMD(DYS-HAC2)#α. Parental riDMD myoblasts: negative control for dystrophin (exons 5–7); healthy myoblasts: positive control. Western blot for dystrophin (427 kDa) in differentiated riDMD and riDMD(DYS-HAC2)#α myoblasts; human muscle and differentiated human inducible myogenic cells (Maffioletti et al, 2015) were used as positive control; differentiated DMD-inducible myogenic cells (Maffioletti et al, 2015) were used as negative control and myosin heavy chain (MyHC) as normaliser (40 μg of proteins loaded for all samples but human muscle, which had 30 μg loaded). Immunofluorescence images showing in vitro muscle differentiation of riDMD myoblasts (negative control), riDMD(DYS-HAC2)#α and healthy donor myoblasts (positive control). Red: MyHC; green: dystrophin; blue: Hoechst. Scale bar: 50 μm. Source data are available online for this figure. Source Data for Figure 2 [emmm201607284-sup-0005-SDataFig2.jpg] Download figure Download PowerPoint hTERT and Bmi1 expression prevents replicative senescence of human mesoangioblasts The experimental work described so far provides proof-of principle evidence of HAC transfer into human dystrophic myoblasts. However, although myoblast transplantation appears to be a promising therapeutic strategy for localised forms of muscular dystrophy (Perie et al, 2014), it appears of limited value for widespread muscle disorders such as DMD, where their modest migration potential is a major hurdle (Tedesco et al, 2010; Skuk & Tremblay, 2014). To overcome this limitation, we extended the DYS-HAC platform to human mesoangioblasts, which can be delivered systemically via the arterial circulation and have been recently assessed in a phase I/II clinical trial based upon allogeneic transplantation (Cossu et al, 2015). Post-natal human mesoangioblasts are considered to be the in vitro progeny of a subset of alkaline phosphatase (ALP)-positive skeletal muscle pericytes (Dellavalle et al, 2007). Firstly, human mesoangioblasts were isolated from muscle biopsies of three different healthy subjects (H#1, H#2 and H#3) according to the standard protocols (Tonlorenzi et al, 2007). To exclude myoblast cross-contamination, cells were FACS-purified as ALP-positive/CD56-negative (see Materials and Methods). After a short in vitro expansion, H#1, #H2 and H#3 human mesoangioblasts were co-transduced with LOX-TERT-IRESTK and LOX-CWBmi1 lentiviral vectors. As an additional control, cells were transduced with a LOX-GFP-IRESTK (Fig EV2A). Phase contrast microscopy revealed that hTERT + Bmi1 transduced polyclonal populations (Fig 3A, upper row, right images) showed a similar morphology to their control (CTR) counterparts (Fig 3A, upper row, left images). One polyclonal population (hTERT + Bmi1 H#3) was then cloned by limiting dilution and three hTERT + Bmi1 clones were selected for further analysis (namely H#3A, H#3B and H#3C; Fig 3A, lower row). PCR analyses performed on genomic DNA of clonal and polyclonal populations confirmed the presence of hTERT and Bmi1 transgenes (Fig 3B). Transcription of both transgenes was then confirmed by RT–PCR (Fig 3C) and quantitative real-time RT–PCR analyses (Fig 3D). Click here to expand this figure. Figure EV2. Characterisation of immortalised mesoangioblasts Phase contrast (upper row) and fluorescence (lower row) of GFP H#1 and H#2 polyclonal populations and of GFP #B5 clone (from GFP H#3 polyclonal population). Scale bar: 100 μm. Western blot showing Bmi1 expression for hTERT + Bmi1 polyclonal populations (hTERT + Bmi1 H#1 and hTERT + Bmi1 H#2) and untransduced parental population (H#1 and H#2). Gapdh: normaliser. Population doubling curves (PD = logN/log2; N = number of initially plate
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