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TISSUE-SPECIFIC STEM CELLS

Isolation and Characterization of Mesoangioblasts from Facioscapulohumeral Muscular Dystrophy Muscle Biopsies

^aDepartment of Neurosciences, Catholic University School of Medicine "A. Gemelli," Rome, Italy;
^bFondazione Don Carlo Gnocchi, Rome, Italy;
^cStem Cells Research Center, San Raffaele Hospital, Milan, Italy

Key Words. Muscle stem cells • Mesoangioblasts • Myoblasts • Facioscapulohumeral muscular dystrophy

Correspondence: Roberta Morosetti, M.D., Department of Neurosciences, Catholic University School of Medicine, Largo A. Gemelli 8, 00168 Rome, Italy. Telephone: 39-06-30154303; Fax: 39-06-35501909; e-mail: rmorosetti{at}rm.unicatt.it; Massimiliano Mirabella, M.D., Ph.D., Department of Neurosciences, Catholic University School of Medicine, Largo A. Gemelli 8, 00168 Rome, Italy. Telephone: 39-06-30154303; Fax: 39-06-35501909; e-mail: mirabella{at}rm.unicatt.it

Received on June 14, 2007; accepted for publication on August 23, 2007.

First published online in STEM CELLS EXPRESS  August 30, 2007.

	ABSTRACT

Top Abstract Introduction Materials and Methods Results Discussion Disclosure of Potential... Acknowledgments References

 
Facioscapulohumeral muscular dystrophy (FSHD) is the third most frequent inherited muscle disease. Because in FSHD patients the coexistence of affected and unaffected muscles is common, myoblasts expanded from unaffected FSHD muscles have been proposed as suitable tools for autologous cell transplantation. Mesoangioblasts are a new class of adult stem cells of mesodermal origin, potentially useful for the treatment of primitive myopathies of different etiology. Here, we report the isolation and characterization of mesoangioblasts from FSHD muscle biopsies and describe morphology, proliferation, and differentiation abilities of both mesoangioblasts and myoblasts derived from various affected and unaffected muscles of nine representative FSHD patients. We demonstrate that mesoangioblasts can be efficiently isolated from FSHD muscle biopsies and expanded to an amount of cells necessary to transplant into an adult patient. Proliferating mesoangioblasts from all muscles examined did not differ from controls in terms of morphology, phenotype, proliferation rate, or clonogenicity. However, their differentiation ability into skeletal muscle was variably impaired, and this defect correlated with the overall disease severity and the degree of histopathologic abnormalities of the muscle of origin. A remarkable differentiation defect was observed in mesoangioblasts from all mildly to severely affected FSHD muscles, whereas mesoangioblasts from morphologically normal muscles showed no myogenic differentiation block. Our study could open the way to cell therapy for FSHD patients to limit muscle damage in vivo through the use of autologous mesoangioblasts capable of reaching damaged muscles and engrafting into them, without requiring immune suppression or genetic correction in vitro.

Disclosure of potential conflicts of interest is found at the end of this article.

	INTRODUCTION

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Facioscapulohumeral muscular dystrophy (FSHD; Online Mendelian Inheritance in Man 158900 [OMIM] ) is an autosomal dominant disease with an estimated prevalence of 1:20,000, corresponding to the third most frequent form of inherited muscle disease, following Duchenne muscular dystrophy (DMD) and myotonic dystrophy. In FSHD, a high variability is observed with regard to age at onset, severity, and pattern of muscle involvement, both between and within families, and the rate of disease progression appears to be unpredictable in a single affected individual [1, 2].

The gene responsible for the disease has been mapped on the long arm of chromosome 4 (region 4q35). This region carries a long stretch of 3.3-kilobase (kb) KpnI repeat units (the D4Z4), and the clinical features of FSHD develop when a deletion reduces the number of the repeats below a critical threshold. The 4q35 telomeric probe p13E-11 detects BlnI-resistant EcoRI polymorphic fragments larger than 40 kb in healthy individuals and smaller fragments (10–40 kb in size, consisting of 1–10 KpnI units) in both sporadic and familial cases [3].

To date, there is a general agreement that FSHD pathogenesis is determined by an alteration in the epigenetic control over a broad chromosomal region, whereby an impairment in the structure of the D4Z4 repeated region translates into a change in chromatin structure, as evidenced by the hypomethylated state of the D4Z4 alleles in FSHD patients [4]. Overexpression of 4q35 proximal genes such as ANT1, FRG1, and FRG2, the last one being the closest gene to D4Z4, have been found in FSHD muscle [5, 6]. Recently, it has been shown that transgenic mice overexpressing the FRG1 gene in muscle tissue develop a muscular dystrophy resembling FSHD [7]. As FRG1 might play a role in pre-mRNA splicing, FRG1 overexpression and consequent alteration of the splicing pattern of specific genes are proposed to underlie muscle degeneration in FSHD. Nevertheless, very little is yet known about the molecular mechanisms activated in FSHD muscles that induce progressive muscle degeneration. In FSHD patients, it is quite common to observe the coexistence of affected and apparently normal muscles. In previous studies, myoblasts obtained from FSHD-affected muscles displayed reduced proliferation ability and impaired differentiation [8]. On the contrary, cells expanded from unaffected FSHD muscles showed normal growth and regenerating ability and were proposed as suitable tools in view of clinical trials with autologous cell transplantation [9]. Although myoblast-based cell therapy protocols have failed for several yet unknown reasons, these observations are relevant in view of the possible alternative use of newly identified myogenic mesodermal stem cells localized in the perivascular niche of muscle tissue, called mesoangioblasts. These vessel-associated cells have been shown to restore muscle morphology and function in animal models of muscular dystrophy when injected intra-arterially [10, 11], a feature that makes them an ideal candidate for muscle reconstitution therapy via cell transplantation. Human mesoangioblasts might be especially useful for muscle regenerative stem cell therapy in diseases such as FSHD, where a very selective muscle involvement occurs.

We recently demonstrated that human adult mesoangioblasts can easily be isolated with high efficiency from diagnostic muscle biopsies of patients with inflammatory myopathies and expanded in vitro to an amount of cells suitable for a potential in vivo treatment [12].

Here, we (a) report the isolation and characterization of mesoangioblasts from FSHD muscle biopsies and (b) describe the main biological properties, in terms of morphology, proliferation, and differentiation abilities, of both mesoangioblasts and myoblasts obtained from three different muscle compartments of patients of both sexes, representative of a large proportion of FSHD patients.

Our data demonstrate that it is possible to grow a transplantable number of mesoangioblasts from small specimens obtained even through a needle biopsy. This could open the way to cell therapy for FSHD patients by using autologous mesoangioblasts, which do not require immune suppression or genetic correction in vitro, thus exploiting their ability to reach damaged muscle and engraft into them.

	MATERIALS AND METHODS

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Patients
Muscle biopsies were performed after informed consent at the Neuroscience Department of Catholic University. We used fresh and fresh-frozen muscles obtained from nine patients with genetically confirmed diagnosis of FSHD (24–67 years old; mean, 39.89 ± 12.42) and three individuals (48–84 years old; mean, 64 ± 18.33) deemed free of neuromuscular disease after all diagnostic tests were performed. In our FSHD patients, the size of the pathogenic EcoRI fragment ranged between 17 and 33 kb, indicating the presence of at least three KpnI repeats. No patients had infantile onset or carried the very short fragments of 10–13 kb associated with the most severe cases of the disease. Age at biopsy ranged between 24 and 67 years. Muscle biopsy was performed on deltoid muscle in two patients, on quadriceps in six (five were needle biopsies), and on paraspinal lumbar muscles (during orthopedic surgery) in one. None of the patients was ever treated with steroids or immunosuppressive therapy. This research was approved by the ethical committee of Catholic University. In each patient (Pt), the overall clinical severity was assessed using the Clinical Severity Scale (CSS) [13], assigning a score 2 when the disease involved only facial and scapular muscles and a score of 3 (mildly affected), 3.5 (moderately affected), or 4–5 (severely affected) when pelvic and lower limb muscles were involved. To assess whether the biopsied muscle was affected and to evaluate the severity of its involvement, we used clinical, histopathologic, and muscle imaging criteria. Magnetic resonance imaging (MRI) study was performed using a 1.5 Tesla magnetic resonance scanner. Axial images were obtained from psoas to distal foot muscles, along with coronal slices of the lumbar and pelvic muscles.

Cell and Organ Cultures
Tissue fragments of approximately 2 mm³, including intramuscular vessels and surrounding mesenchymal tissue, were plated, and explants were cultured for a period ranging from 10 to 21 days as previously described [10, 12, 14]. Cells were maintained in culture for several passages. At every passage, cells were counted, and viability was determined by trypan blue exclusion. Details are provided in the supplemental online Methods.

At passage 7, mesoangioblasts were dissociated to single cells and plated for clonogenic assays as described elsewhere [12, 14].

Characterization of Human Mesoangioblasts from FSHD by FACS
Previous studies have shown that human mesoangioblasts are homogenously positive for CD44, CD13, and CD49b and homogeneously negative for CD34, CD133, and CD45, with a low percentage of cells being vascular endothelial growth factor (VEGF)-RII (KDR)-positive [12, 14]. Details are provided in the supplemental online Methods.

Alkaline Phosphatase Staining
Human mesoangioblasts express the pericyte marker alkaline phosphatase (ALP) [12, 14]. Mesoangioblasts were fixed with 4% paraformaldehyde (PF) and incubated with 0.1 mg/ml of naphthol AS-MX phosphate, 0.5% N,N-dimethylformamide, 2 mM MgCl₂, and 0.6 mg/ml fast blue BB salt in 0.1 M Tris-HCl, pH 8.5 (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com).

Cell Cycle Analysis and Growth Curve of FSHD Mesoangioblasts
Cells at 70% confluence were collected, and nuclei were isolated and stained as previously described [15]. Cell-cycle phases were determined with propidium iodide, enabling calculation of the percentages of cells in G0/G1, S, and G2/M. Flow cytometric DNA ploidy analysis was performed by analyzing a minimum of 10,000 nuclei using an Epics XL flow cytometer (Beckman Coulter, Fullerton, CA, http://www.beckmancoulter.com), and cell cycle analysis was performed using the ModFit software (Verity Software House, Topsham, ME, http://www.vsh.com).

For growth curve, exponentially growing cells were seeded at appropriate concentrations to prevent confluence for the duration of the experiment. Viable cells were counted after 24, 48, and 72 hours to determine the amount of cell proliferation.

Skeletal Muscle Differentiation of FSHD Mesoangioblasts
Human mesoangioblasts differentiate down the skeletal muscle pathway under a variety of conditions, with the highest efficiency when exposed to medium conditioned by normal human myoblasts. In particular, normal human myoblasts at 90% confluence were exposed to growth medium optimized for their expansion containing 5% bovine serum albumin (BSA) (without serum). After 4 days, myoblast-conditioned medium was collected and filtered. Mesoangioblasts were cultured in conditioned medium for 4 days and then exposed for 7 days to differentiation medium with 1% BSA (without serum). At each time point, cells were fixed for immunocytochemistry or harvested by centrifugation for protein extraction. Fusion index was expressed as number of myonuclei per number of total nuclei, visualized by Hoechst 33258 staining (Molecular Probes Inc., Eugene, OR, http://probes.invitrogen.com).

Gene Expression Profiling of FSHD Mesoangioblasts
Mesoangioblasts from Pt 4, Pt 7, and two controls were harvested before and after 4 days of exposure to myoblast-conditioned medium. Total RNA was isolated using the RNeasy RNA isolation kit (Qiagen, Hilden, Germany, http://www.qiagen.com). cDNA synthesis was performed according to the Affymetrix standard protocol. GeneChip staining was performed following protocols for the Hg-Focus gene chip format (Affymetrix, Santa Clara, CA, http://www.affymetrix.com). The eight gene chips were processed in a single labeling and hybridization session. Data analyses were performed using BRB-ArrayTools [16–18]. Raw expression data visualization and quality control were performed using software packages dChip (http://biosun1.harvard.edu) and Bioconductor project (http://www.bioconductor.org) [19]. Gene level summaries were computed using the log scale robust multiarray analysis procedure [20]. The similarity between gene expression patterns was measured by computing the correlation distances for each pair of samples based on standardized log-transformed expression values across all of the genes. Using a matrix of one minus Pearson's correlation (centered) distance measurements from the complete pairwise comparison of all the cell populations, a multidimensional scaling method (MDS) was used to display the overall similarity in gene expression profiles [16–18].

Quantitative Real-Time Polymerase Chain Reaction
Quantitative real-time polymerase chain reaction (q-RT-PCR) for Id1, Id4, HoxC6, and EBF2 in mesoangioblasts from four FSHD patients and three controls. Details are provided in the supplemental online Methods.

Primary Muscle Cultures of FSHD
Fragments from the same nine FSHD muscle biopsies were also cultured to obtain primary muscle cultures from satellite cells using the explantation re-explantation method [21, 22]. Cells were maintained in a replicative state for approximately seven passages using a medium containing 15% serum and a cocktail of growth factors [22]. To induce myogenic differentiation, cells were shifted to a medium with 5% serum and without growth factors (differentiation medium) for 5–7 days. Fusion index was expressed as number of myonuclei per number of total nuclei, visualized by Hoechst 33258 staining (Molecular Probes). Cells were harvested at different time points for Western blot analysis or fixed with 4% PF for immunocytochemistry.

Telomere Restriction Fragment Length Assay
Mean length of terminal restriction fragment was measured using the TeloTAGGG telomere length assay kit (Roche Molecular Biochemical, Indianapolis, IN). Details are provided in the supplemental online Methods.

Bromodeoxyuridine Incorporation
Cells were cultured with 20 µM bromodeoxyuridine (BrdU) (Sigma-Aldrich) for 24 hours and fixed with 4% PF. DNA samples were denatured for 30 minutes at 37°C with a phosphate-buffered saline-Triton X and HCl solution and rinsed with 0.1 M sodium borate. Double immunofluorescence for muscle-specific myosin heavy chain and BrdU was performed. As positive control, we used proliferating myoblasts.

Clonogenic Assays and Growth Curves of FSHD Myoblasts
At passage 3, cells at 70% confluence were dissociated to single cells and cloned by limited dilutions in 48-well dishes. When colonies had reached a minimum of 40 cells, clones were counted.

For growth curve, proliferating myoblasts were seeded at appropriate concentrations to prevent confluence for the duration of the experiment. Viable cells were counted after 24, 48, and 72 hours to determine cell proliferation.

Immunocytochemistry
Cells were cultured on gelatin-coated optical quality plastic µ-Dishes (Integrated DNA Technologies, Munich, Germany, http://www.idtdna.com/Home/Home.aspx) and fixed in 4% PF for 15 minutes. The following primary antibodies were used: polyclonal anti-myosin (Sigma-Aldrich), monoclonal anti-BrdU, and monoclonal anti-desmin (both from Chemicon, Temecula, CA, http://www.chemicon.com). Detection of immunocomplexes was performed using the appropriate secondary antibodies conjugated with either Texas Red or fluorescein isothiocyanate (Jackson Immunoresearch Laboratories, West Grove, PA, http://www.jacksonimmuno.com). Hoechst 33258 (Molecular Probes) staining was used to visualize cell nuclei. Samples were analyzed using a TCS SP5 laser scanning confocal microscope (Leica, Wetzlar, Germany, http://www.leica.com).

Western Blot Analysis
Cells were harvested and homogenized in lysis buffer. Equal amounts of protein (30 µg) were separated by SDS-polyacrylamide gel electrophoresis and blotted onto nitrocellulose membranes (Schleicher and Schuell, Relliehausen, Germany, www.schleicher-schuell.com/bioscience). Blots were incubated with one of the following antibodies: monoclonal anti-skeletal Myosin fast (MY-32; Sigma-Aldrich); monoclonal anti-Myogenin (FD-5; Santa Cruz Biotechnology Inc., Santa Cruz, CA, http://www.scbt.com); monoclonal anti-p21, polyclonal anti-myocite enhancer factor-2C (MEF2C), polyclonal anti-p38 and anti-phospho-p38, polyclonal anti-AKT, and anti-phospho-AKT (all from Cell Signaling Technology, Beverly, MA, http://www.cellsignal.com); monoclonal anti-BrdU (Chemicon); and monoclonal anti-β-actin (Sigma-Aldrich). After incubation with the appropriate horseradish peroxidase-conjugated secondary antibody, blots were visualized using enhanced chemiluminescence (Amersham Biosciences, Piscataway, NJ, http://www.amersham.com) and quantified by densitometry.

Statistical Analysis
All data reported were expressed as means ± SD of triplicates of a representative experiment performed at least three times. One-way analysis of variance was used to compare differences between groups. Statistical significance was set at p .05.

	RESULTS

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Clinical Severity and Muscle Characterization
We included in this study nine FSHD patients different in terms of severity of clinical involvement and size of the EcoRI fragment. Table 1 reports the clinical and molecular data of the patients studied and the relevant information on each biopsied muscle. Seven of the nine FSHD patients had clinical evidence of pelvic and lower limb muscle involvement with a CSS score 3 [13]. Conversely, only two patients (Pt 2 and Pt 8) had clinical manifestations of the disease restricted to the facial and scapular districts (CSS 1 and 1.5, respectively, indicating mild scapular involvement).

View larger version (141K): [in this window] [in a new window]   Figure 1. Morphological features of FSHD muscles. (A): Histopathologic abnormalities (mild endomisial fibrosis, scattered necrotic fibers, and foci of inflammatory infiltration) found in Pt 4. Scale bar = 50 µm. (B–G): Right quadriceps magnetic resonance imaging (MRI) axial images of the four Pts who received the muscle biopsy at this level (*, site of biopsy on the vastus lateralis, which appeared normal in Pt 2 and mildly, moderately, and severely affected and replaced by fatty-fibrous tissue in Pts 1, 3, and 6, respectively). (B): Pt 1; (C): Pt 2; (D): Pt 3; (E): Pt 6; (F): Pt 8; (G): Pt 9. (H–I): Axial and coronal MRI images, respectively, of Pt 5 (the white arrows indicate the approximate site of biopsy). Abbreviation: Pt, patient.

View larger version (86K): [in this window] [in a new window]   Figure 2. Phase contrast; cell morphology. (A): Using a medium optimized for human mesoangioblasts, from the organ culture on, we obtained a double cell population: refractive triangular adherent cells and round, loosely adherent/floating cells. (B): A fragment from the same muscle biopsy was cultured to obtain primary muscle cultures from satellite cells. Scale bars = 100 µm.

Clonogenic Potential, Cell Cycle, and Phenotypic Characteristics Do Not Differ Among FSHD Mesoangioblasts
Cells were dissociated to single-cell suspension, and after 14 days of culture at low density, clones appeared in 11.4% ± 1.8% of wells with the same double morphology of the starting population (Fig. 5C). By replating the clones with a clonal density, they were able to give rise to new clones. We examined the cell cycle characteristics of mesoangioblasts isolated from the seven FSHD muscle biopsies, and the cell cycle distribution was similar for all mesoangioblasts examined, regardless of the muscle impairment (not shown).

The phenotypic characterization was performed by analyzing expression of the surface markers recently identified for human mesoangioblasts [12, 14]. Cells from all patients, analyzed at various passages, were strongly positive for CD44 and CD13, positive for CD49b, and homogeneously negative for CD34, CD133, and CD45, with a low percentage of cells being VEGF-RII (KDR)-positive (supplemental online Fig. 2). In addition, all FSHD mesoangioblasts were positive for ALP, a marker identifying human adult mesoangioblasts as the in vitro progeny of pericytes (supplemental online Fig. 3).

FSHD Mesoangioblasts Differentiate into Skeletal Muscle
Human mesoangioblasts differentiate into skeletal muscle under a variety of conditions, with the highest efficiency when exposed to human normal myoblast-conditioned medium. Therefore, we cultured mesoangioblasts from nine FSHD biopsies (seven with myopathic signs and two with normal morphology) and three normal controls in conditioned growth medium for 4 days and subsequently in differentiation medium for 7 days. Interestingly, despite their normal characteristics (morphology, proliferation rate, clonogenicity, and cell cycle distribution), FSHD mesoangioblasts differentiated into skeletal muscle to a different extent, which was apparently correlated with the overall disease severity (as evaluated by the CSS) and the degree of involvement of the muscle of origin (as evaluated by clinical, MRI, and histopathological examination). Mesoangioblasts from the six morphologically abnormal muscles obtained from the more severely affected patients (CSS 3.5–4.5) were able to fuse only into spare myosin-positive myotubes (fusion index 0.15 ± 0.11). Moreover, these myotubes were short, and most of them contained no more than two nuclei. The paraspinal muscle biopsy of Pt 5 gave rise to a higher percentage of cells able to fuse into myosin-positive thin and mainly binucleated myotubes (fusion index, 0.3 ± 0.05), whereas from the apparently spared quadriceps of Pt 2 and Pt 8, 60%–70% of mesoangioblasts differentiated into multinucleated myosin-positive myotubes (fusion index, 0.7 ± 0.08), with no significant difference from normal control mesoangioblasts (Fig. 3A). Western blot analysis of differentiating mesoangioblasts showed, in all samples, an upregulation of Myogenin (not shown), MEF2C, and Myosin absent in undifferentiated mesoangioblasts and characteristic of myogenic differentiation, with the highest levels observed in the morphologically normal biopsies (Fig. 3B; supplemental online Fig. 4).

View larger version (37K): [in this window] [in a new window]   Figure 3. Skeletal muscle differentiation of FSHD mesoangioblasts. (A): Immunofluorescence for myosin. Mesoangioblasts from unaffected muscles spontaneously fused into multinucleated, fully differentiated myotubes (60%–70%); cells obtained from affected muscles were able to fuse into spare (10%–15%) myosin-positive myotubes. Scale bar = 50 µm. Each experiment was performed in duplicate at least two times for each patient. (B): For Western blot analysis, cells were harvested in growth medium (0) after 4 days in myoblast-conditioned medium and 7 days in differentiation medium. A marked upregulation of myosin could already be observed at day 4 in conditioned medium in mesoangioblasts from unaffected muscle, whereas only a mild upregulation of myosin was visible at day 7 in cells from severely affected muscle. Abbreviation: FSHD, facioscapulohumeral muscular dystrophy; MEF2C, anti-myocite enhancer factor-2C.

View larger version (21K): [in this window] [in a new window]   Figure 4. FSHD mesoangioblasts constitutively express high levels of the helix-loop-helix proteins ID1, ID2, ID3, and ID4 that are upregulated after 4 days of exposure to myoblast-conditioned medium. T0, basal level; T4, day 4 in conditioned medium. Abbreviations: Pt, patient; rma, robust multi-array average.

All FSHD myoblasts fully differentiated into multinucleated myosin-positive myotubes; however, the size of FSHD myotubes was often larger than that of controls (average nuclei per myotube, 66.44 ± 51.33 vs. 17.69 ± 9.70). In particular, eight of nine FSHD cultures gave hypertrophic terminally differentiated multinucleated myotubes, as indicated by complete absence of BrdU-positive nuclei (Fig. 6A, Fig. 6B). Moreover, we explored different pathways involved in skeletal muscle differentiation, and no clear-cut differences were observed between FSHD myoblasts from affected and unaffected muscles (Fig. 6C).

View larger version (38K): [in this window] [in a new window]   Figure 5. Morphology, proliferation ability, clonogenicity, and senescence of FSHD myoblasts. (A): Immunofluorescence for desmin. Myoblasts from FSHD biopsies display a higher number of vacuoles in the cytoplasm compared with controls; however, no abnormalities in cytoplasm shape or necrotic cells could be observed. Scale bar = 30 µm (left panel), 5 µm (right panels). A representative culture of FSHD myoblasts is shown. (B): Growth curve. Cell growth was assessed after 24, 48, and 72 hours. Results are expressed as absolute counts. Bars represent the mean ± SD of triplicate samples of one representative experiment out of three performed in seven FSHD muscle cultures compared with three normal controls. (C): Clonogenic assays of FSHD myoblasts and mesoangioblasts. The clonogenic ability of FSHD myoblasts was significantly lower than that of controls (p .05). By contrast, no significant difference between FSHD and control mesoangioblasts was observed (p .05). (D): Hybridization of digested myoblast DNA to digoxigenin-labeled probe specific for telomeric repeats. Measurements of the telomeric smear in normal control (passage 6), a Pt severely affected (Pt 1) (passage 6), and one Pt with normal muscle biopsy (Pt 2) (passage 5). Terminal restriction fragments from the unaffected Pt ranged in size from approximately 4.5 to 10 kbp, which is shorter than affected muscle and normal control smear (5–20 kbp). Ct-DNA high and Ct-DNA low values were supplied by TeloTAGGG telomere length assay kit. Abbreviations: Ct, control; FSHD, facioscapulohumeral muscular dystrophy; kbp, kilobase pairs; Pt, patient.

View larger version (76K): [in this window] [in a new window]   Figure 6. All FSHD myoblasts fully differentiated into multinucleated myosin-positive myotubes. (A): Immunofluorescence for myosin in myotubes obtained from myoblasts from affected (Pt 3, Pt 4, and Pt 9) and apparently unaffected (Pt 8) muscles. Scale bar = 100 µm. (B): Immunofluorescence for myosin and bromodeoxyuridine (BrdU) from affected (Pt 1) and apparently unaffected (Pt 2) muscles. Virtually all myotubes were terminally differentiated with no BrdU-positive nuclei. Red-fluorescent, Texas Red-labeled nuclei are visible within myoblasts. Six representative cultures of FSHD muscle cells are shown. Scale bar = 100 µm. (C): Western blot analysis of FSHD myoblasts during differentiation. No differences between FSHD myoblasts (from both affected and unaffected muscles) and normal controls were detected in the pathways involved in skeletal muscle differentiation. A representative experiment is shown. (D): Myotubes derived from differentiating mesoangioblasts (left) and from satellite cells (right) of the same muscle biopsy cultured simultaneously in identical differentiating conditions show completely different morphology. Scale bar = 100 µm. Abbreviations: FSHD, facioscapulohumeral muscular dystrophy; MEF2C, anti-myocite enhancer factor-2C; Pt, patient.

Normal human mesoangioblasts can spontaneously differentiate into skeletal muscle when cultured in low serum on Matrigel-coated (BD Biosciences, San Diego, http://www.bdbiosciences.com) dishes (20%–40%) [14]. However, to obtain a higher differentiation efficiency (60%–70%), they need to be exposed to a "myogenic environment" either by coculture with murine myoblasts or by exposure to a medium previously conditioned by human myoblasts for 4 days [12]. To verify whether FSHD myoblasts from severely affected muscles were capable of producing the myogenic factors required to induce the muscle commitment of normal human mesoangioblasts, we challenged normal mesoangioblasts with growth medium conditioned by FSHD myoblasts. Indeed, growth medium conditioned by FSHD myoblasts was able to trigger the myogenic commitment of these stem cells similarly to the one obtained from normal myoblasts (not shown).

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion Disclosure of Potential... Acknowledgments References

 
In this study, we demonstrated that human mesoangioblasts can be efficiently isolated from muscle biopsies of FSHD patients and that it is possible to expand them even from a needle biopsy specimen to obtain a reasonable number of cells to transplant into an adult patient. Proliferating mesoangioblasts from nine muscle biopsies examined did not differ from normal controls in terms of morphology, phenotypic characteristics, proliferation rate, or clonogenicity. However, even though proliferating FSHD mesoangioblasts appeared morphologically normal, their ability to differentiate into skeletal muscle was partially impaired, and this differentiation defect correlated with the overall disease severity and the degree of histopathologic alterations of the examined muscle of origin. In fact, only mesoangioblasts from "normal" muscles of the less severely affected FSHD patients differentiated normally. By contrast, the differentiation ability of cells from the paraspinal muscle with minimal myopathic changes and normal clinical and MRI findings was partially impaired, whereas a remarkable defect of differentiation was observed in mesoangioblasts from all mildly to severely affected deltoid and quadriceps muscles.

We show that ID4, which is known to inhibit the DNA-binding of E47 homodimers as well as E47/MyoD heterodimers [24], was stably overexpressed in comparison with controls and upregulated after exposure to myoblast-conditioned medium (which is crucial for the skeletal muscle commitment) in FSHD mesoangioblasts. These observations raise the possibility that inhibitory pathways of mesoangioblast differentiation may be activated in the course of chronic myopathies.

We also examined primary muscle cells derived from the same muscle biopsies used to isolate mesoangioblasts and, except for a modest vacuolization, they did not show a significant difference in cytoplasmic shape or other structural abnormalities compared with controls. By contrast, all FSHD myoblast cultures displayed decreased proliferation and clonogenic abilities compared with normal controls. In addition, the cells from one unaffected muscle also exhibited reduced life span, as shown by telomere length. In spite of the abnormalities described above, all myoblasts from FSHD patients fully differentiated into multinucleated myotubes, with no differences in the modulation of the myogenic pathways explored.

Defects in skeletal muscle differentiation have been proposed by different studies showing a dysregulation of genes involved in myogenesis (such as MyoD or MEF2), cell differentiation, and cell-cycle control in FSHD muscle [25, 26], as well as morphological alterations such as irregular cytoplasmic shape and a "vacuolar/necrotic" phenotype in cultured FSHD myoblasts [8]. Vilquin et al. [9] tested growth and differentiation abilities of myoblasts obtained from muscles apparently spared by the disease and showed no differences between myoblasts of FSHD patients and healthy donors, thus suggesting that autologous transplantation of myoblasts expanded from unaffected muscles could improve the impaired repair/regeneration capability of the affected muscles.

Even if a primary dysregulatory effect of myogenesis takes place in FSHD, it is conceivable that the disease process may not affect to the same degree satellite cells and a different myogenic stem cell population such as mesoangioblasts, which are vessel-associated stem cells. Unlike satellite cells, which represent the main source for adult muscle repair and regeneration after injury, in normal muscle mesoangioblasts probably contribute to a lesser degree to generation of new muscle fibers. However, mesoangioblasts appear to be greatly activated in response to specific chemoattractant factors and during muscle damage and inflammation, although their quantitative contribution to muscle regeneration in physiological and pathological conditions remains to be fully elucidated. According to our results, FSHD mesoangioblasts showed an impairment of skeletal muscle differentiation even when minimal histopathologic features were the only evidence of disease in the muscle of origin; moreover, we obtained a very low percentage of differentiating cells from all mildly to severely affected deltoid and quadriceps muscles regardless any difference in the age of patients, EcoRI fragment size, clinical severity, or degree of muscle pathology. It is noteworthy that upregulation of vascular-associated genes has been recently reported as characteristic of the molecular pathogenesis of FSHD muscle [27], leading, together with the high incidence of retinal vasculopathy reported in FSHD patients, to the hypothesis that abnormalities within the skeletal muscle vascular system may contribute to the disease process [28]. Therefore, it could be hypothesized that an abnormal microvascular system of FSHD muscles may also cause abnormal activation or variable functional impairment of vessel-associated stem cells, such as mesoangioblasts.

Our results on satellite-derived myoblasts do not show a clear-cut distinction between affected and unaffected muscles in terms of morphological abnormalities and differentiation abilities. Our data also indicate that in FSHD it is truly difficult and may be overly simplistic to categorize as "uninvolved" by the disease a given muscle district, even by complementing clinical examination with morphological investigation of muscle by MRI and histology. Although limited in number, our cohort spans a representative selection of FSHD patients, and apparent discrepancies observed between our results and those of previous studies may be correlated to the different characteristics of patients from whom myoblasts were obtained (i.e., age and size of the pathogenic EcoRI fragment). Genotype-phenotype correlation studies on large cohorts of FSHD patients have shown that with the increase of the EcoRI fragment size (i.e., number of KpnI repeats), the risk of developing a lower limb muscle involvement progressively decreases as the number of nonpenetrant gene carriers rises [13, 29, 30]. It is not currently possible to conclude whether apparently unaffected muscles can indeed be considered normal, and more important, it remains to be fully clarified whether primary or secondary defects in the regenerative capability of muscle involving satellite cells play a pathogenic role in the disease.

Our study, however, demonstrates that apparently unaffected muscles are a valuable source of mesoangioblasts that can be isolated and extensively grown in culture and induced to differentiate into myotubes that apparently do not reproduce the morphological abnormalities observed in primary myoblast cultures obtained from the same muscles. We show that mesoangioblasts may also easily be isolated from affected muscles, but the myogenic differentiation ability of these cells is variably but significantly impaired, although proliferation and life span do not differ from normal controls.

The clinical use of stem cells to correct gene defects in devastating hereditary muscle diseases such as DMD requires in vitro manipulation of defective cells or, probably more easily, the use of healthy donor cells associated with chronic immune suppression to prevent rejection. As proposed in acquired degenerative muscle diseases such as inclusion-body myositis [12] or in peculiar genetic muscle diseases, such as FSHD, in which a variable percentage of muscle groups may be spared lifelong, a new strategy of treatment with myogenic stem cells may be envisaged. In fact, with the goal being to halt the disease progression and limit muscle damage, rather than to obtain a specific gene correction, it is conceivable to use in vitro expanded autologous stem cells capable of engrafting into damaged muscles and enhancing their regenerative activity. To this purpose, mesoangioblasts offer the great advantage of possible systemic delivery, opening the way to future studies using these cells expanded in vitro from unaffected muscles of patients with FSHD. It is obviously compelling to expand our knowledge on the biological characteristics of mesoangioblasts isolated from muscles of patients with different age, clinical severity and EcoRI fragment size, and on various possible pharmacological and biomolecular strategies to increase their survival, migration, and myogenic capabilities, as well as to revert a differentiation block when detected. Equally important is the analysis of putative environmental cues related to the specific molecular defect of the disease acting in the pathological muscle milieu, to minimize the detrimental effects on the survival and the effectiveness of the transplanted mesoangioblasts.

	DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST

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The authors indicate no potential conflicts of interest.

	ACKNOWLEDGMENTS

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This work was supported by Grant Linea D3.2 from Catholic University and by grants from the Italian Ministries of Health and University and Research. We thank Manuela Papacci and Gabriella Proietti for valuable technical assistance. R.M. and M.M. contributed equally to this work.

	REFERENCES

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1. Padberg GW. Facioscapulohumeral disease [M.D. thesis] Leiden, The Netherlands: Leiden University,1982;.

2. Lunt PW, Harper PS. Genetic counselling in facioscapulohumeral muscular dystrophy. J Med Genet 1991;28:655–664.[Abstract/Free Full Text]

3. Tupler R, Gabellini D. Molecular basis of facioscapulohumeral muscular dystrophy. Cell Mol Life Sci 2004;61:557–566.[CrossRef][Medline]

4. van Overveld PG, Enthoven L, Ricci E et al. Variable hypomethylation of D4Z4 in facioscapulohumeral muscular dystrophy. Ann Neurol 2005;58:569–576.[CrossRef][Medline]

5. Gabellini D, Green MR, Tupler R. Inappropriate gene activation in FSHD: A repressor complex binds a chromosomal repeat deleted in dystrophic muscle. Cell 2002;110:339–348.[CrossRef][Medline]

6. Rijkers T, Deidda G, van Koningsbruggen S et al. FRG2, an FSHD candidate gene, is transcriptionally up-regulated in differentiating primary myoblast cultures of FSHD patients. J Med Genet 2004;41:826–836.[Abstract/Free Full Text]

7. Gabellini D, D'Antona G, Moggio M et al. Facioscapulohumeral muscular dystrophy in mice overexpressing FRG1. Nature 2006;439:973–977.[Medline]

8. Winokur ST, Barrett K, Martin JH et al. Facioscapulohumeral muscular dystrophy (FSHD) myoblasts demonstrate increased susceptibility to oxidative stress. Neuromuscul Disord 2003;13:322–333.[CrossRef][Medline]

9. Vilquin JT, Marolleau JP, Sacconi S et al. Normal growth and regenerating ability of myoblasts from unaffected muscles of facioscapulohumeral muscular dystrophy patients. Gene Ther 2005;12:1651–1662.[CrossRef][Medline]

10. Sampaolesi M, Torrente Y, Innocenzi A et al. Cell therapy of alpha-sarcoglycan null dystrophic mice through intra-arterial delivery of mesoangioblasts. Science 2003;301:487–492.[Abstract/Free Full Text]

11. Sampaolesi M, Blot S, D'Antona G et al. Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs. Nature 2006;444:574–579.[CrossRef][Medline]

12. Morosetti R, Mirabella M, Gliubizzi C et al. MyoD expression restores defective myogenic differentiation of human mesoangioblasts from inclusion-body myositis muscle. Proc Natl Acad Sci U S A 2006;103:16995–17000.[Abstract/Free Full Text]

13. Ricci E, Galluzzi G, Deidda G et al. Progress in the molecular diagnosis of facioscapulohumeral muscular dystrophy and correlation between the number of KpnI repeats at the 4q35 locus and clinical phenotype. Ann Neurol 1999;45:751–757.[CrossRef][Medline]

14. Dellavalle A, Sampaolesi M, Tonlorenzi R et al. Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat Cell Biol 2007;9:255–267.[CrossRef][Medline]

15. Broccolini A, Gidaro T, Morosetti R et al. Neprilysin participates in skeletal muscle regeneration and is accumulated in abnormal muscle fibres of inclusion body myositis. J Neurochem 2006;96:777–789.[CrossRef][Medline]

16. Simon R, Korn E, McShane L et al. Design and Analysis of DNA Microarray Investigations. New York: Springer-Verlag,2003;.

17. Korn EL, Troenndle JF, McShane LM et al. Controlling the number of false discoveries: Application to high-dimensional genomic data. J Stat Plan Inference 2004;124:379–398.[CrossRef]

18. Simon R, Lam A. BRB-ArrayTools User Guide, version 3.2. Bethesda, MD: Biometric Research Branch, National Cancer Institute, NIH,2004;.

19. Gentleman RC, Carey VJ, Bates DM et al. Bioconductor: Open software development for computational biology and bioinformatics. Genome Biol 2004;5:R80.[CrossRef][Medline]

20. Irizarry RA, Bolstad BM, Collin F et al. Summaries of Affymetrix GeneChip probe level data. Nucleic Acids Res 2003;31:e15.[Abstract/Free Full Text]

21. Askanas V, Engel WK. Cultured normal and genetically abnormal human muscle. In: Rowland LP, Di Mauro S, eds. Handbook of Clinical Neurology.Amsterdam, The Netherlands: Elsevier Science Publisher,1992;85–116.

22. Broccolini A, Ricci E, Pescatori M et al. Insulin-like growth factor I in inclusion-body myositis and human muscle cultures. J Neuropathol Exp Neurol 2004;63:650–659.[Medline]

23. Chen Y-W, Zhao P, Borup R et al. Expression profiling in the muscular dystrophies: Identification of novel aspects of molecular pathophysiology. J Cell Biol 2000;151:1321–1336.[Abstract/Free Full Text]

24. Riechmann V, van Cruchten I, Sablitzky F. The expression pattern of Id4, a novel dominant negative helix-loop-helix protein, is distinct from Id1, Id2 and Id3. Nucleic Acids Res 1994;22:749–755.[Abstract/Free Full Text]

25. Winokur ST, Chen YW, Masny PS et al. Expression profiling of FSHD muscle supports a defect in specific stages of myogenic differentiation. Hum Mol Genet 2003;12:2895–2907.[Abstract/Free Full Text]

26. Celegato B, Capitanio D, Pescatori M et al. Parallel protein and transcript profiles of FSHD patient muscles correlate to the D4Z4 arrangement and reveal a common impairment of slow to fast fibre differentiation and a general deregulation of MyoD-dependent genes. Proteomics 2006;6:5303–5321.[CrossRef][Medline]

27. Osborne RJ, Welle S, Venance SL et al. Expression profile of FSHD supports a link between retinal vasculopathy and muscular dystrophy. Neurology 2007;68:569–577.[Abstract/Free Full Text]

28. Fitzsimons RB, Gurwin EB, Bird AC. Retinal vascular abnormalities in facioscapulohumeral muscular dystrophy. A general association with genetic and therapeutic implications. Brain 1987;110:631–648.[Abstract/Free Full Text]

29. Lunt PW, Jardine PE, Koch MC et al. Correlation between fragment size at D4F104S1 and age at onset or at wheelchair use, with a possible generational effect, accounts for much phenotypic variation in 4q35-facioscapulohumeral muscular dystrophy (FSHD). Hum Mol Genet 1995;4:951–958.[Abstract/Free Full Text]

30. Tawil R, Forrester J, Griggs RC et al. Evidence for anticipation and association of deletion size with severity in facioscapulohumeral muscular dystrophy. The FSH-DY Group. Ann Neurol 1996;39:744–748.[CrossRef][Medline]

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