Cell Therapy

original article

© The American Society of Gene & Cell Therapy

Human Skeletal Muscle–derived CD133+ Cells Form Functional Satellite Cells After Intramuscular Transplantation in Immunodeficient Host Mice Jinhong Meng1, Soyon Chun1, Rowan Asfahani1, Hanns Lochmüller2, Francesco Muntoni1 and Jennifer Morgan1 1 The Dubowitz Neuromuscular Centre, UCL Institute of Child Health, London, UK; 2Institute of Genetic Medicine, Newcastle University, International Centre for Life, Central Parkway, Newcastle upon Tyne, UK

Stem cell therapy is a promising strategy for treatment of muscular dystrophies. In addition to muscle fiber formation, reconstitution of functional stem cell pool by donor cells is vital for long-term treatment. We show here that some CD133+ cells within human muscle are located underneath the basal lamina of muscle fibers, in the position of the muscle satellite cell. Cultured hCD133+ cells are heterogeneous and multipotent, capable of forming myotubes and reserve satellite cells in vitro. They contribute to extensive muscle regeneration and satellite cell formation following intramuscular transplantation into irradiated and cryodamaged tibialis anterior muscles of immunodeficient Rag2-/γ chain-/ C5-mice. Some donor-derived satellite cells expressed the myogenic regulatory factor MyoD, indicating that they were activated. In addition, when transplanted host muscles were reinjured, there was significantly more newly-regenerated muscle fibers of donor origin in treated than in control, nonreinjured muscles, indicating that hCD133+ cells had given rise to functional muscle stem cells, which were able to activate in response to injury and contribute to a further round of muscle regeneration. Our findings provide new evidence for the location and characterization of hCD133+ cells, and highlight that these cells are highly suitable for future clinical application. Received 18 October 2013; accepted 16 February 2014; advance online publication 18 March 2014. doi:10.1038/mt.2014.26

INTRODUCTION

Determining the optimal muscle stem cells to repair and regenerate skeletal muscle is essential for effective treatment of muscular dystrophies. A suitable stem cell for therapeutic purposes should not only survive and make muscle fibers following transplantation, but also participate in the reconstitution of a functional stem cell pool, able to repair, and regenerate muscle fibers throughout life. The classical skeletal muscle stem cell is the satellite cell, located in its niche between the sarcolemma and basal lamina

of muscle fibers. Satellite cells are a heterogeneous population1 that are responsible for growth, repair, and regeneration of muscle; mouse satellite cells are also capable of self-renewal, to generate functional satellite cells.2 Other stem cells, including pericytes, mesoangioblasts, and CD133+ cells that are present within skeletal muscle have been shown to contribute to muscle regeneration in animal models.3–8 Among the human muscle–derived stem cells so far investigated, only myoblasts9,10 and CD133+ cells11 give rise to satellite cells after intramuscular transplantation into immunodeficient mice. Although there is evidence that satellite cells formed by injected myoblasts are functional,9,10 the therapeutic potential of human myoblasts is limited by their poor transplantation efficiency (reviewed in ref. 12). Thus, the human skeletal muscle–derived CD133+ (hCD133+) cell becomes a promising candidate stem cell type

for future cell therapy due to its relatively higher transplantation efficiency than human myoblasts,11 ability to be systemically delivered to skeletal muscle and transducibility by lentiviral vectors.3,11 However, the anatomical location of this cell population within human muscle is unknown, and although hCD133+ cells transplanted into mouse muscles were shown to form Pax7+ cells located in the satellite cell position, the functionality of these donor-derived satellite cells was not tested.11 This is important, as cells from other sources, e.g., the bone marrow, can enter the satellite cell position, but are not functional.13 Donor-derived satellite cells must be functional to ensure their long-term therapeutic role within the host muscle. To understand more about the location, phenotype, and long term therapeutic potential of hCD133+ cells, we investigated their anatomical location on transverse sections of human skeletal muscle, examined their characteristics and myogenic properties in vitro and their contribution to muscle regeneration within injured muscles of Rag2-/γ chain-/C5-mice. We provide the first evidence of the anatomical position of CD133+ cells within human muscle. In addition, we show that hCD133+ cells effectively participate in muscle regeneration and give rise to functional satellite cells after intramuscular transplantation into host mice, evidence that they could be exploited for treating muscular dystrophy.

Correspondence: Jennifer Morgan, The Dubowitz Neuromuscular Centre, UCL Institute of Child Health, 30 Guilford Street, London, WC1N 1EH, UK. E-mail: [email protected]

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Human CD133+ Cells form Functional Satellite Cells

RESULTS CD133+ cells are present within normal and Duchenne muscular dystrophy human muscles, either inside or outside the muscle fiber basal lamina

We found no CD133+ cells in muscle sections from two control patients (Table 1; patients 1 and 2) which might be due to their

extremely low incidence within normal muscle.3 However, in muscle sections taken from neonatal muscle (from two 18-dayold nondystrophic control patients (Table 1; patients 6, 7)), we detected CD133+ cells located at the periphery of the muscle fiber, underneath the basal lamina, coexpressing the satellite cell marker Pax7 (Figure 1a–d), suggesting that a subset of satellite cells in

Table 1 List of muscle biopsies used for analysis Muscle ID

1

Age

Muscle type

10 years 10 months

Not known

Diagnosis

Application

Normal (minimal changes on muscle biopsy)

IF for CD133

Summary of results

Negative

2

3 years 5 months

Quadriceps

Mitochondrial myopathy

IF for CD133

Negative

3

2 years 10 months

Quadriceps

DMD

IF for CD133

4

6 years 10 months

Quadriceps

DMD

IF for CD133

Negative CD133+ cells inside

5

7 years 5 months

Quadriceps

DMD

IF for CD133

6

18 days

Not known

Immature muscle (no abnormalities on muscle biopsy)

IF for CD133

7

18 days

Not known

Immature muscle (Abnormal M-oxidation fatty acids, minimal changes on muscle biopsy)

IF for CD133

8

14 years

Paraspinal

Adolescent idiopathic scoliosis (control)

Isolation of CD133+ cells, bulk culture

N/A

9

14 years 11 months

Paraspinal

Adolescent idiopathic scoliosis (control)

Isolation of CD133+ cells, bulk culture

N/A

10

15 years 3 months

Paraspinal

Adolescent idiopathic scoliosis (control)

Isolation of CD133+ cells, bulk culture

N/A

11

15 years 8 months

Paraspinal

Adolescent idiopathic scoliosis (control)

Isolation of CD133+ cells, bulk culture, transplantation

N/A

and outside basal lamina CD133+ cells inside

and outside basal lamina CD133+ cells in the

satellite cell position

CD133+ cells in the satellite cell position

DMD, Duchenne muscular dystrophy.

a

b

c

c

d

e

i

k

m

j

l

n

f

h g

Figure 1 CD133+ cells in human muscle sections. Sections were stained with antibodies to CD133 (green), Pax7 (red), and pan-laminin (magenta in b and d, red in e, j, l, and n), nuclei were counter stained with DAPI (blue). (a,b) Sections of 18-day-old normal human muscle. (c,d) Enlarged images of square c and d within a and b, respectively. CD133 (green) is present on Pax7+ (red) satellite cells (a and c) located underneath the basal lamina of muscle fibers (b and d) in developing human muscles. Bar = 10 µm. (e) CD133+ cells within a section of DMD human muscle. Square f, g, and h highlight three individual CD133+ cells (green) which were located either underneath (i and j) or outside the basal lamina (red, k– n). (i–n) Corresponding enlarged images of squares f–h. (i, k, m) show staining with green (CD133) and blue (DAPI), j, l, and n depict staining with red (laminin), green (CD133), and blue (DAPI), showing the location of each CD133+ cell. MF, muscle fiber. Bar = 5 µm. DAPI, 4′,6-diamidino-2-phenylindole; DMD, Duchenne muscular dystrophy.

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Human CD133+ Cells form Functional Satellite Cells

neonatal human muscle express CD133. In addition, we detected CD133+ cells in muscle sections of two out of three Duchenne muscular dystrophy (DMD) patients (Table 1; patients 3, 4, and 5), located either underneath the basal lamina of myofibers (satellite cell position, Figure 1e,f,i,j) or in an interstitial position, outside muscle fibers (Figure 1e,g,h,k–n).

CD133+ cells isolated from human muscle give rise to cells of different mesenchymal lineages in vitro The number of CD133+ cells in each cell preparation (n = 4, Table  1, patients 8–11) was too low to count immediately after magnetic-activated cell sorting. Colonies of CD133+ cells appeared after 5–10 days in culture, their morphology being similar in the three different proliferation media (see Supplementary Figure S1a–c). Characterization was performed on proliferating cells of two cell preparations (Table 1; patients 8 and 9) at mean population doubling (mpd) 9.45–13.08. Immunostaining showed

a

b

c

d

© The American Society of Gene & Cell Therapy

that the progeny of bulk cultured CD133+ cells contained satellite cells/myoblasts (Pax7+, Myf5+, MyoD+, desmin+, CD56+, and M-cadherin+), pericytes (ALP+, PDGFRβ+, NG2+, and αSMA+) and mesenchymal stem cells (CD49b; see Supplementary Figure S2). Fluorescence-activated cell sorting (FACS) analysis of the cultured CD133+ cells showed that 74.9% expressed the myoblast marker CD56, 0.022% expressed CD34, 0.126% expressed the endothelial cell lineage marker CD31, 2.64% expressed the pericyte marker ALP, 15.8% expressed PDGFR-β, and 10% expressed CD146. Other mesenchymal lineage markers—CD90, CD44, and Stro-1—were expressed by 36.4, 99.4, and 92.4% of cells, respectively (see Supplementary Figure S3).

hCD133+ cells are myogenic in vitro All preparations of CD133+ cells differentiated into multinucleated myotubes in vitro. One typical cell preparation (Table 1; patient 9) was induced to differentiate for 7 days and quantified at mpd 8.29. As shown in Figure 2a–d, the fusion index of this cell preparation is 42.46 ± 3.01% (mean ± SEM), determined by the percentage of nuclei within myosin+ myotubes/total nuclei; myotubes are defined as containing at least three nuclei. Pax7+ cells (6.2 ± 1.02%, mean ± SEM) were also present between the myotubes, evidence of formation of reserve satellite cells by CD133+ cells during differentiation (Figure 2a–d). Dystrophin, a marker of mature myotubes, was expressed in differentiated myotubes derived from CD133+ cells (Figure 2e–h). hCD133+ cells contribute to muscle regeneration hCD133+ cells maintained in three different proliferating media

e

f

g

h

Figure 2 Myogenicity of hCD133+ cells in vitro. (a–d) Formation of myotubes (expressing myosin, red, c) and reserve satellite cells (Pax7, green, b) by hCD133+ cells which have been induced to differentiate for 7 days in culture. Nuclei were counterstained with DAPI (blue, a). Bar = 25 µm. (e–h) Mature myotubes derived from hCD133+ cells express dystrophin in vitro. Myotubes were double stained with antibodies to myosin (red, g) and dystrophin (green, f). Nuclei were counterstained with DAPI (blue, e). Bar = 25 µm. DAPI, 4′,6-diamidino-2-phenylindole.

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were transplanted at mpd 7.15–8.29 into tibialis anterior (TA) muscles of Rag2-/γ chain-/C5-mice. Four weeks after transplantation, human lamin A/C+ nuclei and human spectrin+ myofibers were detected in all grafted muscles. The number of human lamin A/C+ nuclei, human spectrin+ fibers, and human spectrin+ fibers containing human lamin A/C+ nuclei were quantified (Figure 3a–f) and compared among three groups using one-way analysis of variance followed by Tukey’s multiple comparison test. Overall, hCD133+ cells expanded in all three media contributed to muscle regeneration (Figure 3a–f; Table 2). There were no significant differences in the number of donor cells (human lamin A/C+ nuclei) among the three groups (P = 0.0856). There were differences in the number of human spectrin+ fibers (P = 0.0472) among the three groups. A more rigorous definition of a fiber of human origin is one that contains two human-specific markers, e.g., human spectrin and a human lamin A/C+ nucleus (defined here as S+L fibers).14–16 Using this criterion to identify fibers of donor origin, we found that there were also significant differences of S+L fibers among the three groups (P = 0.0123). Further, Tukey’s multiple comparison test confirms that there were significantly more human spectrin+ fibers in muscles of group 1 than in group 2, and significantly more S+L fibers in group 3 than group 2, but there was no difference in the number of S+L fibers between group 1 and 3. In summary, hCD133+ cells survive in our in vivo mouse model and contribute to robust muscle regeneration after they had www.moleculartherapy.org  vol. 22 no. 5 may 2014

© The American Society of Gene & Cell Therapy

Human CD133+ Cells form Functional Satellite Cells

a

c

d hLaminA/C Spectrin S+L

800 600 400 200

hLaminA/C

f

Spectrin S+L

4,000 3,000 2,000 1,000 1,000

No. positive cells/fibres

4,000 3,000 2,000 1,000 1,000

No. positive cells/fibres

800 600 400 200

1

2

3

4

5

6

7

8

1

No. positive cells/fibres

hLaminA/C Spectrin S+L

800 600 400 200

2

3

4

5

1

6

g

j

h

k

l

Low mpd cells

500

1 month

400

3 month

300 200 100

2

3

4

5

6

Muscle ID

Muscle ID

Muscle ID

i

4,000 3,000 2,000 1,000 1,000

0

0

0

No. positive cells/fibres

No. positive cells/fibres

b

e

High mpd cells

300

1 month 3 month

200

100

*

*

0

0 hLaminA/C

Spectrin

S+L

hLaminA/C

Spectrin

S+L

Figure 3 hCD133+ cells contribute to robust muscle regeneration after intramuscular transplantation into irradiated and cryodamaged TA muscles of Rag2-/γ chain-/C5-mice. Muscle sections were stained with human lamin A/C and human spectrin (both green). Nuclei were counterstained with DAPI (blue). Bar = 25 µm. (a–f) Comparison of in vivo myogenic properties of hCD133+ cells maintained in medium 1 (a, b), medium 2 (c, d), and medium 3 (e, f). a, c, e shows representative images of the transplanted muscle; b, d, f are graphs showing the number of human lamin A/C+ nuclei, human spectrin+ fibers, and human spectrin+ fibers containing at least one human lamin A/C+ nucleus (S+L) in each transplanted muscle. Bar = 25 µm. (g–l) Comparison of the contribution to muscle regeneration of hCD133+ cells, which were grafted at low (low mpd cells, g–i) and high population doublings (high mpd cells, j–l) 1 month (g, j) and 3 months (h, k) after transplantation. (i, l) Comparison of the number of human lamin A/C+ nuclei, human spectrin+ fibers, and human spectrin+ fibers containing at least one human lamin A/C+ nucleus (S+L) 1 and 3 months after transplantation with (i) low mpd cells or (l) high mpd cells. Bar = 25 µm. DAPI, 4′,6-diamidino-2-phenylindole; mpd, mean population doubling; TA, tibialis anterior.

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Human CD133+ Cells form Functional Satellite Cells

Table 2 Comparison of transplantation efficiency of hCD133+ cells grown in different media Marker

Number

Medium 1 (n = 6)

Medium 2 (n = 6)

Medium 3 (n = 8)

One-way ANOVA

Range

170–1,019

24–3,456

593–2,272

P = 0.0856

Mean ± SE

698.1 ± 115

1,724 ± 519.5

1,416 ± 310.6

51–749

0–105

78–338

44.83 ± 14.90

177.0 ± 36.39

hLamin A/C hSpectrin

Range Mean ± SE

S+L

Range Mean ± SE

276.8 ± 82.42 27–168 93.25 ± 20.16

0–55 26.67 ± 8.269

60–249

P = 0.0472* P = 0.0123*

126.7 ± 26.68

ANOVA, analysis of variance. *Significant difference

Table 3 Comparison of transplantation efficiency of low or high mpd cells at different time point Cells

Time after grafting

hLamin A/C

hSpectrin

S+L

Low mpd cells (mean ± SE)

1 month (n = 6)

372.3 ± 65.64

211.3 ± 38.54

111.8 ± 12.41

3 months (n = 4)

204.5 ± 26.28

166.8 ± 4.008

107.8 ± 6.921

P value (Mann–Whitney test) High mpd cells (mean ± SE)

0.1143

0.4762

1 month (n = 5)

174.8 ± 40.20

195.2 ± 54.66

95.00 ± 30.63

3 months (n = 5)

110.4 ± 24.39

39.6 ± 17.06

20.4 ± 9.416

P value (Mann–Whitney test)

0.2222

0.0159*

0.7619

0.0317*

mpd, mean population doubling. *Significant difference

been cultured in all three media, but cells maintained in media 1 and 3 contribute to significantly more muscle regeneration than those expanded in medium 2. Subsequent experiments were therefore performed on cells that had been grown in media 1 or 3.

hCD133+ cells that had undergone greater expansion in vitro contribute less to muscle regeneration in vivo The expansion of mouse and human myoblasts in vitro significantly reduces their contribution to muscle regeneration in mouse models in vivo.14,17 To identify whether this is also the case with hCD133+ cells, we transplanted them at low (7.15–8.29) or high (18.19) mpd into our in vivo experimental model, and compared their contribution to muscle regeneration 1 or 3 months after transplantation. Cells transplanted at the lower mpd gave rise to a similar number of human nuclei, human spectrin+ fibers, and human spectrin+ fibers containing a human nucleus at 1 and 3 months after grafting. In contrast, although the same cells transplanted at the higher mpd gave rise to similar numbers of human lamin A/C+ nuclei 1 and 3 months after grafting, the number of human spectrin+ fibers and of S+L fibers were both significantly higher at 1 than at 3 months after transplantation (Figure 3g–l and Table 3). Donor CD133+ cells give rise to Pax7+ cells located both underneath and outside the basal lamina of muscle fibers Knowing that hCD133+ cells can contribute to muscle regeneration after intramuscular transplantation, the next question we asked was whether they could also form satellite cells. To identify satellite cells of donor origin, we stained transverse sections of group 1 muscles with antibodies to human lamin A/C and the satellite cell marker Pax7 (Figure 4A). The number of Pax7+ cells (which include cells of both human and mouse origin) in representative 1012

sections of each muscle (n = 8) was 40.5 ± 12.18 (mean ± SEM); the number of human lamin A/C+/Pax7+ cells was 38.25 ± 12.10 (mean ± SEM), which accounts for 91.8 ± 3.5% of total Pax7+ cells, showing that the majority of Pax7+ cells were of human origin (Figure 4A). This is not surprising, as the host muscle had been irradiated, which leads to a significant loss of host satellite cells.18 To determine whether donor-derived Pax7+ cells were in the satellite cell position, triple labeling of human lamin A/C, Pax7, and laminin was performed and the sections were observed under the confocal microscope. Donor-derived Pax7+ cells were located both in the satellite cell position (underneath the basal lamina of myofibers; Figure 4B) and outside the basal lamina (Figure 4C). There were 11.5 ± 4.93 and 26.75 ± 7.42 human lamin A/C+/ Pax7+ cells per representative transverse section (equivalent to 24.45 ± 4.64 and 75.55 ± 4.64% of total Pax7+ cells of human origin) located inside and outside basal lamina, respectively, indicating that although donor cells gave rise to Pax7+ cells in vivo, only a quarter of these cells are bone fide satellite cells, the remainder being myoblasts located outside the satellite cell niche.

Donor-derived satellite cells are functional in vivo For treatment of muscular dystrophies, an ideal stem cell should not only survive and differentiate into muscle fibers to rescue the acute muscle injury, but it should also reconstitute the stem cell pool to fulfill long-term therapeutic benefit to the host muscle. However, a cell of donor origin in the satellite cell position is not necessarily functional.13 A subset of donor-derived satellite cells express MyoD, a marker of activated satellite cells To test the functionality of the hCD133+ cell-derived satellite cells, we firstly examined whether satellite cells of donor origin were able to express MyoD, a marker of either an activated satellite www.moleculartherapy.org  vol. 22 no. 5 may 2014

© The American Society of Gene & Cell Therapy

a

b

c

Figure 4 Formation of satellite cells by hCD133+ cells after intramuscular transplantation into irradiated and cryodamaged TA muscles of Rag2-/γ chain-/C5-mice. (A) Double staining of human lamin A/C (red, b) and Pax7 (green, c) on muscle sections that had been transplanted with hCD133+ cells 1 month previously. Nuclei were counterstained with DAPI (blue, a). d shows the merged image of the three channels. Human lamin A/C+ Pax7+ cells (white arrow) were present within the muscle. Bar = 10 µm. (B, C) Multichannel staining of human lamin A/C (red, d), Pax7 (green, c), and pan-laminin (cyan, b) on muscle sections, which had been transplanted with hCD133+ cells. Donor-derived satellite cells (human lamin A/C+ Pax7+) were present both underneath (laminin+, cyan, upper panel, B) and outside the basal lamina of myofibers (laminin+, cyan, lower panel, C). Nuclei were counterstained with DAPI (blue, a). Bar = 10 µm. DAPI, 4′,6-diamidino-2-phenylindole; TA, tibialis anterior.

cell, or its proliferating progeny.19 Multichannel immunostaining (Figure 5A) of human lamin A/C, human spectrin, human MyoD, and pan-laminin on sections of mouse muscles that had been transplanted with hCD133+ cells revealed that a subset of Molecular Therapy  vol. 22 no. 5 may 2014

Human CD133+ Cells form Functional Satellite Cells

human lamin A/C+ cells (red), are located outside the muscle fiber sarcolemma (red, human spectrin+ fiber), underneath basal lamina (cyan, pan-laminin+), colocalized with 4′,6-diamidino2-phenylindole (blue), were also expressing MyoD (green), a marker of activated satellite cells (white arrow; Figure 5A). In addition, donor nuclei (human lamin A/C+) expressing human MyoD (green, yellow arrow) but inside the sarcolemma of the myofiber (human spectrin+) were also seen in the same section, suggesting that these were recently formed myonuclei of human origin within the regenerated muscle fibers.

Cells of hCD133+ origin within grafted muscles formed functional muscle stem cells The fact that a proportion of donor-derived satellite cells expressed MyoD indicates that they are activated, rather than quiescent, and are therefore to some extent functional. To determine whether the donor-derived satellite cells were indeed fully functional, we tested their response to an imposed injury to the muscle. The finding of newly regenerated muscle fibers of donor origin a week after the grafted, regenerated muscle had been injured by notexin is evidence that the transplanted donor cells had given rise to functional satellite cells in vivo.2 To our surprise, we found small fibers with high human neonatal myosin expression (i.e., recently regenerated myofibers)20 in transplanted muscles that had not been injured by notexin. Expression of neonatal myosin suggests that these irradiated, cryodamaged muscles were not fully reinnervated.21 We also found that the number of recently regenerated fibers closely correlated with the transplantation efficiency, indicated by the number of human spectrin+ fibers (Figure 5Ba and b) quantified within the same muscle section. To rule out the effect of the initial transplantation efficiency on the number of recently-regenerated fibers of donor origin, we normalized the number of the recently regenerated fibers to the number of human spectrin+ fibers, and compared the muscle treated with notexin to its contralateral, nontreated muscle, analyzed by paired t-test. The ratio of the number of recently regenerated muscle fibers/number of total donor-derived muscle fibers (human spectrin+) was significantly higher in notexin treated than in nontreated muscles (P = 0.0008), evidence of functional satellite cells of donor origin which gave rise to newly regenerated muscle fibers following injury.

DISCUSSION

The human skeletal muscle–derived CD133+ cell is a promising muscle stem cell type for cell therapy of muscular dystrophies such as DMD.3,11 These cells can be expanded in vitro, generating large numbers of cells for transplantation and are myogenic in vitro. Our findings confirm previous work showing that hCD133+

cells contribute to robust muscle regeneration after intramuscular injection in an immunodeficient mouse model.11 Importantly, we provide novel evidence that the transplanted cells functionally reconstituted the satellite cell pool, suggesting their potential for long-term treatment of muscle diseases. CD133 is a transmembrane protein which is highly expressed in stem cells such as hematopoietic stem cells,22 neural stem cells,23 cancer stem cells,24 and very small embryonic like stem cells.25,26 Although CD133+ cells have been prepared 1013

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Human CD133+ Cells form Functional Satellite Cells

a

b

a hspec+/nmy− s.b nmy+

7

5

6 Mouse ID

Mouse ID

6

4 3

c

5 4 3

2

2

1

1 0

200

400

hspec+/nmy− s.b nmy+

7

600

% Small bright hNMY fibres

b

0

100

200

300

No. positive Fibers

No. positive Fibers

−NTX

+NTX

400

500

80 60 40 20 0 −NTX

+NTX Treatment

c

Figure 5 Donor-derived satellite cells are functional in vivo. (A) Multichannel immunostaining of human lamin A/C (red, b and g), human spectrin (red, b and g), MyoD (green, c and h), and pan-laminin (cyan, d and i) on sections of irradiated and cryodamaged muscles that had been transplanted with hCD133+ cells 1 month previously. The white arrow indicates a donor-derived satellite cell, verified by its expression of human lamin A/C (red), located outside the muscle fiber sarcolemma (spectrin+, red) but inside the basal lamina (laminin+, cyan), expressing MyoD (green), a marker of activated satellite cells. Yellow arrow points to a donor-derived myonucleus, expressing human lamin A/C (red), but located inside the muscle fiber (spectrin+, red), also expressing MyoD (green). (a–e) Bar = 25 µm, i–j are higher magnificent images taken from the same area as a–e. Bar = 5 µm. (B) Quantification of the number of human spectrin+ fibers (blue + orange bars) and the number of recently regenerated fibers defined as small, bright neonatal myosin+ fibers (s.b. NM+, orange bars) in each individual grafted, regenerated muscles that had either not been injured by notexin (a, −NTX) or injected with notexin 7 days previously (b, +NTX). The ratio of neonatal myosin+ fibers versus human spectrin+ fibers (normalized) in both groups of muscles was compared using paired t-test (c). There were a significant difference between notexin treated and nontreated groups (P = 0.0008). (C) Immunostaining of human lamin A/C (red, b and f), human spectrin (red, b and f), and human neonatal myosin (green, c and g) in transplanted muscles treated with notexin (upper panel, a–d) or control, noninjured (lower panel e–h). Nuclei were counterstained with DAPI (blue, a and e). Bar = 200 µm. DAPI, 4′,6-diamidino-2-phenylindole; NM, neonatal myosin; s.b. NM+, small, bright neonatal myosin+; NTX, notexin.

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from human skeletal muscle,3,11 the origin of these cells has not been identified. We showed that CD133+ is expressed by a subset of satellite cells (expressing Pax7) in human skeletal muscles. In neonatal normal muscles and postnatal DMD muscles, CD133+ cells were present in the satellite cell position as well as in the interstitium. In contrast, in nondystrophic adult human muscle, no CD133+ cells were seen in the sections examined, although CD133+ cells can be isolated from such muscles. The finding of CD133+ cells within very young normal and DMD muscles and their rarity or absence in muscles derived from two juvenile controls (Table 1; patients 1 and 2) suggests that they are implicated in muscle growth and regeneration. Following their expansion in culture, hCD133+ cells give rise to cells expressing different lineage markers (see Supplementary Figures S2 and S3), suggesting that they are either a mixed population of cells within skeletal muscle, or that they are multipotent stem cells, able to give rise to several mesenchymal lineages in culture. They express Pax7, which identifies quiescent satellite cells, in their niche between the basal lamina and sarcolemma of muscle fibers.27–29 In injured muscle, satellite cells become activated, proliferate and may leave their niche, to repair neighboring damaged muscle fibers. After proliferation as Pax7/MyoD-expressing myoblasts, most cells maintain MyoD but downregulate Pax7 and commit to myogenic differentiation. Other myoblasts maintain Pax7 but downregulate MyoD, withdraw from the cell cycle, and reexpress markers of quiescent satellite cells, including Pax7.31-33 Pax7 therefore identifies both satellite cells and a proportion of their progeny myoblasts. When placed under in vitro conditions that promote myogenic differentiation, hCD133+ cells formed multinucleated myotubes and Pax7+ reserve34–38 cells, evidence of myogenic differentiation and contribution to the satellite cell pool. When transplanted intramuscularly in our mouse model, hCD133+ cells contributed to robust muscle regeneration and also gave rise to functional satellite cells, able to contribute to a further round of muscle regeneration following reinjury of the host muscle (Figures 3–5). However, although hCD133+ cells are promising for therapeutic application in muscular dystrophies, particularly for targeted treatment of key muscles by intramuscular delivery, they have several drawbacks. They are very rare within skeletal muscle of normal adults, representing 1% of the total mononucleated muscle cells.3 They are also very fragile cells; in our hands, they did not survive FACS sorting, necessitating the use of a milder, but possibly less specific method (magnetic-activated cell sorting) to isolate them from skeletal muscle. Expansion of these cells in culture, which would be necessary for clinical application, is challenging. Human blood–derived CD133+ cells are difficult to expand in vitro,39 and the only medium previously shown to support expansion of muscle-derived CD133+ cells is not readily available.3,11 We show that skeletal muscle–derived hCD133+ cells could proliferate in all three different commercially available media that we tested, but that they contributed to muscle regeneration to a significantly greater extent when they had been expanded in M10 and EGM-2 medium than in CD133+ cell proliferation medium. Similarly to myoblasts, prolonged culture of donor hCD133+ cells reduces their effectiveness in vivo. Whether hCD133+ cells prepared from different donors are uniform in Molecular Therapy  vol. 22 no. 5 may 2014

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their phenotype and differentiative potential is also a concern, but we found no significant differences in the myogenic capacity of cells prepared from paraspinal muscles of all four normal donors. In addition, the contribution to muscle regeneration of systemically-delivered hCD133+ cells needs to be confirmed,3 as we failed to obtain any contribution to muscle regeneration from intraarterially delivered hCD133+ cells in our mouse model. This might be due to various factors, including the phenotype of cells used for transplantation, different genetic and immunodeficient background of the recipient mice and the injury models used. As systemic delivery is required for treatment of muscular dystrophies such as DMD, in which muscles are affected body-wide, more efforts are need to explore the mechanism by which donor stem cells transmigrate through the blood vessel and contribute to muscle regeneration in some dystrophic animal models,3,4,7,8 but not others.15 Nonetheless, based on our findings that hCD133+ cells are stem cells present within human skeletal muscle and that, apart from myoblasts, they are the only human stem cell that contributes to a functional muscle stem cell pool after transplantation, it is worthwhile to consider moving to clinical trials with these cells. Ideally, to rule out the potential immune rejection by the recipient, hCD133+ cells could be isolated from the patient, manipulated in vitro to express, e.g., dystrophin or an exon-skipping construct, then transplanted back to the patient. Even if hCD133+ cells are not routinely or efficiently systemically deliverable, they would be useful for treatment of key muscles, such as finger muscles in DMD patients,40 or muscles that are more affected by a muscular dystrophy, e.g., oculopharyngeal muscular dystrophy, in which that eyelid elevator and pharyngeal muscles are primarily affected.41

MATERIALS AND METHODS

Mice were bred and experimental procedures were carried out in the Biological Services Unit, University College London Institute of Child Health, in accordance with the Animals (Scientific Procedures) Act 1986. Experiments were performed under Home Office licence number 70/7086. Experiments were approved by the local University College London ethical committee prior to the licence being granted. Immunofluorescent staining of human muscle sections. Transverse,

cryostat sections of muscle biopsies from control patients (with minimal muscle pathology), young patients (18 days of age, with minimal changes in muscle pathology), and DMD patients were studied. Details of muscle biopsies are listed in Table 1. Seven micrometers cryosections were air dried at room temperature (RT) for 30 minutes before being fixed with 4% paraformaldehyde for 15 minutes at RT. The sections were then washed with phosphatebuffered saline (PBS) three times followed by overnight incubation at 4 °C in mouse anti-CD133 (MiltenyiBiotec, 293C3, 1:100, Surry, UK), Pax7 (DHSB, 1:100, Iowa City, IA), and rabbit antipan-laminin (Sigma, 1:1,000, Dorset, UK) diluted in PBS containing 10% normal goat serum and 0.03% Triton X100. Sections were then washed with PBS and incubated with Alexa594-conjugated goat antimouse IgG2b, Alexa-647-conjugated goat antimouse IgG1, and Alexa-488-conjugated goat antirabbit IgG (H+L) antibodies (Invitrogen, Paisley, UK) at RT for 1 hour. Sections were then mounted with mounting medium (DAKO, Ely, UK) containing 10 µg/ml 4′,6-diamidino-2-phenylindole. Sections stained with secondary antibody only were used as negative control.

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Isolation and maintenance of human muscle CD133+ cells. Biopsies of paraspinal muscle of four adolescent idiopathic scoliosis patients were taken with the patient’s consent. Muscles were cut into 1 mm3 pieces using a scalpel and digested with an enzyme mixture containing collagenase IA-S (Sigma, C9722), II (Sigma, C1764), and IV (Sigma, C1889; 1 mg/ ml each) in 20% fetal bovine serum Dulbecco’s modified eagle’s medium for 45 minutes at 37 °C. The cell suspension was then diluted with PBS and filtered through a 40 µm cell strainer (SLS, 352340, Nottingham, UK) before centrifuged at 300g for 10 minutes at RT. The resulting pellet was resuspended in 10 ml red cell lysis buffer (Cambridge Bioscience, 10089, Cambridge, UK) for 5 minutes at RT, centrifuged at 300g for 10 minutes at RT. The pellet was then incubated with a dead cell removal kit (MiltenyiBiotec, 130-090-101) and the dead cells removed according to the manufacturer’s instructions. The live cells were then centrifuged at 300g for 5 minutes at RT, incubated with antihuman CD133 microbeads (MiltenyiBiotec, 130-050-801), 1:11 at 4 °C for 30 minutes. The CD133+ and CD133− cell populations were separated using LS column (MiltenyiBiotec, 130-042-401) in a magnetic-activated cell sorting system (MiltenyiBiotec). Resulting CD133+ cells were cultured in three different proliferation media and expanded at 37 °C in 5% O2 and 5% CO2 incubator. The media we tested in our study were: (Medium 1): M10 medium: Megacell Dulbecco’s modified eagle’s medium (M3942, Sigma) containing 10% fetal bovine serum, 2 µmol/l glutamine, 1% nonessential amino acids, 0.1 mmol/l β-mercaptoethanol, and 5 ng/ml basic fibroblast growth factor, a medium shown to support the proliferation of human muscle–derived pericytes4,15. (Medium 2): CD133+ cell proliferation medium (Filarete InvestimentiS.p.a., Milan, Italy). (Medium 3): EGM-2 medium (LONZA, CC-3202, Wolverhampton, UK), a medium used to expand human muscle–derived myoendothelial cells.42 Plated CD133+ cells started to proliferate 5–10 days after isolation (see Supplementary Figure S1a–c). Cells were trypsinized when they approached confluence and the cell numbers recorded. For long-term maintenance, cells were plated at a density of 2.5 × 105 cells/75 cm2 flask, with 1 mg/ml collagen I as the substrate and in the media described above. Cells were passaged every 3–4 days, and mean population doublings (mpds) were calculated as previously described.15 Aliquots of cells were frozen at each passage and stored in liquid nitrogen for future studies. In vitro characterization of hCD133+ cells. For the cell phenotype assay,

2 × 104 cells were plated onto 5 µg/ml poly-d-lysine coated eight-well chamber slides (Fisher Scientific, Loughborough, UK) and incubated for 1–3 days before being processed for immunofluorescent staining. Cells were fixed with 4% paraformaldehyde for 15 minutes and incubated with PBS containing 10% normal goat serum/0.03% Triton X100 for 30 minutes. Cells were then incubated with primary antibodies for 1 hour followed by Alexa 488-conjugated goat antimouse or rabbit IgG (H+L; Invitrogen, 1:500) for 1 hour. All staining procedures were performed at RT. Cells stained with secondary antibody only were used as negative control. Primary antibodies used for characterization were: Pax7 (DSHB, 1:100, Iowa City, IA), MyoD (DAKO, M3512, clone 5.8A, 1:50), desmin (DAKO, M0760, clone D33, 1:200), PDGFR-β (AbDSerotec, 1:200, Oxford, UK), CD49b (AbDSerotec, 1:200), M-Cadherin (Nano tools, 0124-100/MCAD21G4, clone 21G4, 1:100, Teningen, Germany), CD34 (MiltenyiBiotec, 1:50), alkaline phosphatase (ALP, Santa Cruz, 1:200, Middlesex, UK), Myf5 (Santa Cruz, sc302, 1:200), α-smooth muscle actin (α-SMA, DAKO, 1:100), NG2 (Millipore, 1:200, Watford, UK). For FACS analysis of cultured hCD133+ cells, proliferating cells at mpd 18.91 were collected by trypsinization and incubated with CD56:PE (MiltenyiBiotec, 1:50), CD34:FITC (MiltenyiBiotec, 1:50), CD31 (AbDSerotec, 1:200), alkaline phosphatase:FITC (ALP, Santa Cruz, 1:200), PDGFR-β (AbDSerotec, 1:50), CD146:FITC (AbDSerotec, 1:50), CD90:PE (AbDSerotec, 1:50), CD44:PE (AbDSerotec, 1:200), and Stro-1 (Millipore, 1:50) for 30 minutes at RT. Cells incubated with nonconjugated antibodies were followed by incubation with rabbit antimouse IgG:RPE

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(AbDSerotec, 1:50) or goat anti mouse IgM: FITC (Millipore, 1:50) for 30 minutes at RT. For each antibody staining, a corresponding isotype control was included for setting up the gate when performing the analysis. Cells were analyzed with BD LSRII FACS machine (BD Biosciences, Oxford, UK). Ten thousand events were collected for each sample. Flowjo 7.2.5 software (Tree Star, Ashland, OR) was used to analyze the results. For myogenic differentiation, CD133+ cells were plated at 5 × 104 cells/ well onto 10 µg/ml laminin (Invitrogen)-coated chamber slides and induced to differentiate in Megacell Dulbecco’s modified eagle’s medium containing 2% fetal bovine serum. Cells were fixed with 4% paraformaldehyde at 7 days after the commencement of differentiation. Immunostaining with antibodies against myosin (MF20, DSHB, 1:100), Pax7, and dystrophin (Fisher Scientific, RB9024P1, 1:200) was performed as described above. Images were acquired using Metamorph software (Molecular Device, Sunnyvale, CA) using a Leica microscope (Leica Microsystems, Milton Keynes, UK). In vivo transplantation and analysis

Intramuscular transplantation of hCD133+ cells. Four- to eight-week-old Rag2-/γ chain-/C5-mice43–45 were used as recipients in this study. Hind legs were irradiated with 18Gy 3 days before cell transplantation.20,46 On the day of transplantation, TA muscles were cryodamaged with three freeze-thaw cycles using a cryoprobe prechilled in liquid nitrogen.14 5 × 105 hCD133+

cells/5 µl culture medium were injected into each TA with a Hamilton syringe. For analyzing donor cell survival, their contribution to regenerated muscle fibers and satellite cells, three groups of recipient mice (both TA muscles of each mouse) were transplanted with CD133+ cells at mpd 7.15–8.29 maintained in three different proliferation media: medium 1 (n = 8), medium 2 (n = 6), and medium 3 (n = 6). Muscles were removed for analysis 1 month after grafting. For comparison of the in vivo muscle regenerative potential of cells at different mpds, CD133+ cells at mpd 7.15–8.29 (early passage cells) or mpd 18.19 (late passage cells) were transplanted to recipient mice as described above. Injected muscles were analyzed at 1 month (n = 6 for early passage cells and n = 5 for late passage cells) and 3 months (n = 4 for early passage cells and n = 5 for late passage cells) after transplantation. For the functional satellite cell assay, CD133+ cells at mpd 7.15–8.29 were transplanted into irradiated and cryodamaged TA muscles of a group of seven mice as described above. Eight weeks (n = 3) and 14 weeks (n = 4) after transplantation, the right TA of the recipient mice was injected with 10 µl of 10 µg/µl notexin, a myotoxin that destroys muscle fibers but spares other resident muscle cells, including satellite cells, to induce a secondary injury to the grafted, regenerated muscle.2 The contralateral, grafted muscle was not injected with notexin (control). The mice were analyzed 1 week after notexin treatment and analyzed as described below. Analysis of muscle sections. Grafted TA muscles were dissected and frozen in isopentane chilled in liquid nitrogen. Eight micrometers transverse cryosections were taken throughout the muscle and stained with antibodies to human spectrin (Vector labs, VP-S283, 1:100, Peterborough, UK), human lamin A/C (Vector labs, VP-L550, 1:100), pan-laminin (Sigma, L9393, 1:1000), Pax7, MyoD, human neonatal myosin (Vector labs, VP-M666, 1:100) followed by corresponding secondary antibodies (Alexa 488-conjugated goat antimouse IgG1, Alexa 594-conjugated goat antimouse IgG2b, Alexa 647, or 594 conjugated goat antirabbit IgG (H+L), etc., Invitrogen). Images were captured with MetaMorph software using a Leica microscope. Four-colour images were acquired using a Zeiss LSM 710 confocal microscope (Carl Zeiss, Cambridge, UK). The number of human lamin A/C+ nuclei, human spectrin+ fibers, human spectrin+ fibers containing human lamin A/C+ nuclei (as a confirmation that the spectrin+ fibers were of donor origin) were counted in representative transverse sections using MetaMorph software to quantify the number of donor cells and their contribution to muscle regeneration. The data were analyzed by one-way analysis of variance or Mann–Whitney test using graphpad prism5 software (GraphPad Software, La Jolla, CA). In the reinjury experiment, the www.moleculartherapy.org  vol. 22 no. 5 may 2014

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number of small fibers that were strongly-expressing human neonatal myosin (newly regenerated muscle fibers) was counted and normalized to the number of human spectrin+ fibers in the same section as a proportion of total fibers of human origin, and the ratio in the right TA (experimental) muscle and the left TA (control) muscle of each mouse was compared and analysed using paired t-test. Ethics. Tissue sampling was approved by the NHS national Research Ethics Service, Hammersmith, and Queen Charlotte’s and Chelsea Research Ethics Committee. Setting up of a rare diseases biological samples bank (biobank) for research to facilitate pharmacological, gene, and cell therapy trials in neuromuscular disorders REC reference number: 06/Q0406/33, in compliance with national guidelines regarding the use of biopsy tissue for research. All patients or their legal guardians gave written informed consent.

SUPPLEMENTARY MATERIAL

Figure S1. Morphology of cultured hCD133+ cells maintained in medium 1 (a), medium 2 (b) and medium 3 (c). Figure S2.  Characterization of bulk cultured hCD133+ cells by immunostaining of various lineage markers (all in green). Figure S3.  Characterization of cultured hCD133+ cells by FACS analysis.

ACKNOWLEDGMENTS The support of the MRC Centre for Neuromuscular Diseases Biobank is gratefully acknowledged. We thank Geraldine Edge for her help in obtaining the muscle biopsies, Lucy Feng, Darren Chambers, and Diana Johnson for their help with providing the human muscle sections for immunostaining. We thank Maximilien Bencze for critical reading of the manuscript. This work was funded by the MRC and the Duchenne Parent Project (Netherlands). J.M. was funded by a Wellcome Trust University Award. F.M. and J.M. are supported by Great Ormond Street Hospital Children’s Charity. J.M., S.C., R.A., and J.M. designed and performed the experiments, analyzed the data, wrote, and edited the paper. F.M. and H.L. read and edited the paper. The authors have no conflicts of interest (both financial and personal).

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