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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www.moleculartherapy.org vol. 22 no. 5, 1008–1017 may 2014
© The American Society of Gene & Cell Therapy
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.
Molecular Therapy vol. 22 no. 5 may 2014
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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.
1010
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.
Molecular Therapy vol. 22 no. 5 may 2014
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© The American Society of Gene & Cell Therapy
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
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© The American Society of Gene & Cell Therapy
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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© The American Society of Gene & Cell Therapy
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
Human CD133+ Cells form Functional Satellite Cells
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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Human CD133+ Cells form Functional Satellite Cells
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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© The American Society of Gene & Cell Therapy
(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
© The American Society of Gene & Cell Therapy
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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