Facioscapulohumeral muscular dystrophy region gene 1 over-expression causes
primary defects of myogenic stem cells.
Running title: Muscle Stem Cell Defects in FRG1 mice
Keywords: muscular dystrophy, muscle stem cells, epigenetics
Alexandros Xynos1, Maria Victoria Neguembor1,2, Roberta Caccia1, Danilo Licastro3, Alessandro
Nonis2, Clelia Di Serio2, Elia Stupka3,4 and Davide Gabellini1
1 Dulbecco Telethon Institute and Division of Regenerative Medicine, San Raffaele Scientific Institute,
2 Università Vita-Salute San Raffaele, Milano, Italy
3 CBM S.c.r.l., Area Science Park, Trieste, Italy
4 Current address: Center for Translational Genomics and Bioinformatics, San Raffaele Scientific
Institute, Milano, Italy
Davide Gabellini, Division of Regenerative Medicine, San Raffaele Scientific Institute, DIBIT 2, 5A3-
44, Via Olgettina 58, 20132 Milano, Italy. E-mail: email@example.com
© 2012. Published by The Company of Biologists Ltd.
Journal of Cell Science
JCS Advance Online Article. Posted on 22 March 2013
Over-expression of FSHD Region Gene 1 (FRG1) in mice, frogs and worms leads to muscular
and vascular abnormalities. Nevertheless, the mechanism that follows FRG1 over-expression and
finally leads to muscular defects is currently unknown. Here, we show that the earliest phenotype
displayed by FRG1 mice is a postnatal muscle-growth defect. Long before the development of
muscular dystrophy, FRG1 mice exhibit also a muscle regeneration impairment. Ex-vivo and in-vivo
experiments revealed that FRG1 over-expression causes myogenic stem-cell activation, proliferative,
clonogenic and differentiation defects. A comparative gene expression profiling of WT and FRG1
muscles from young pre-dystrophic mice identified differentially expressed genes in several gene
categories and networks that could explain the emerging tissue and myogenic stem-cell defects.
Overall, our study provides new insights in the pathways regulated by FRG1 and suggests that muscle-
stem cells defects could contribute to the pathology of FRG1 mice.
Journal of Cell Science
Facioscapulohumeral muscular dystrophy (FSHD, OMIM 158900) is the third most common
muscular dystrophy, exhibits autosomal dominant inheritance and has no cure (Cabianca and Gabellini,
2010). FSHD typically arises with a reduction of facial and shoulder girdle muscle mass. The disease
may extend to abdominal and pelvic girdle muscles impairing the ability to walk. Although FSHD is
primarily a disease of skeletal muscle, up to 75% of FSHD patients also present vascular defects
(Fitzsimons et al., 1987; Osborne et al., 2007; Padberg et al., 1995).
FSHD differs from classical muscular dystrophies for several aspects. While in many
myopathies sarcolemmal disruption is the primary pathogenetic mechanism (Dalkilic and Kunkel,
2003; Durbeej and Campbell, 2002), in FSHD patients there is no evidence for alteration of
sarcolemmal integrity (Orrell et al., 1999) or mitochondrial involvement (Kilmer et al., 1995) and the
mechanism responsible for the disease is currently unclear.
FSHD is associated with reduction in the copy number of a macrosatellite repeat, called D4Z4,
located at the subtelomeric region of chromosome 4 long arm, in 4q35 (Wijmenga et al., 1992). The
number of repeats varies between 11 and 100 in healthy individuals, while FSHD patients carry 1 to 10
repeats (van Deutekom et al., 1993). The contraction of the D4Z4 repeat array causes a
Polycomb/Trithorax epigenetic switch leading to the over-expression of several genes within the FSHD
region (Cabianca et al., 2012; Gabellini et al., 2002; Lemmers et al., 2010; Snider et al., 2009; Snider et
al., 2010). The peculiar nature of the mutation at the basis of FSHD and its complex effect on
chromatin surrounding the 4q35 region makes it highly unlikely that the root cause of the disease can
be attributed to a single gene. Since expression of multiple genes is affected, the molecular
pathogenesis of FSHD has been challenging to untangle, and as yet no therapy is available for FSHD
patients. The two most important FSHD candidate genes are the D4Z4 repeat gene double homeobox 4
(DUX4) (Lemmers et al., 2010; Snider et al., 2009; Snider et al., 2010) and the proximal gene FSHD
Region Gene 1 (FRG1) (Gabellini et al., 2002). Nevertheless, for both DUX4 (Jones et al., 2012;
Tsumagari et al., 2011) and FRG1 (Klooster et al., 2009; Masny et al., 2010) significant controversy
exists regarding their actual over-expression and their role in the disease. For this reason, the potential
role of 4q35 gene over-expression in the disease has been investigated at the functional level. Despite a
lot of effort, transgenic mice over-expressing DUX4 and showing muscle pathology are currently not
available. On the contrary, mice over-expressing FRG1, selectively in the skeletal muscle, display
reduced muscle size and develop a muscular dystrophy resembling FSHD (Gabellini et al., 2006).
Journal of Cell Science
Moreover, studies conducted in X. laevis and C. elegans demonstrated that frg1 is required for normal
muscle development and its over-expression causes muscular defects and vascular abnormalities
correlated with the clinical findings from FSHD patients (Hanel et al., 2009; Liu et al., 2010; Wuebbles
et al., 2009). Collectively, these results suggest that FRG1 is important for muscle function and its
aberrant expression could contribute to the FSHD pathogenesis.
FRG1 is a dynamic nuclear and cytoplasmic shuttling protein that, in skeletal muscle, is also
localized to the sarcomere (Hanel et al., 2011). Interestingly, over-expressed FRG1 is almost
completely nuclear and is localized in nucleoli, Cajal bodies, and actively transcribed chromatin (Sun et
al., 2011; van Koningsbruggen et al., 2004). Although, it has been associated with RNA biology
(Gabellini et al., 2006; Sun et al., 2011; van Koningsbruggen et al., 2004; van Koningsbruggen et al.,
2007), the molecular and cellular mechanism that follows FRG1 over-expression leading to muscular
dystrophy is currently unknown.
To address this point, we monitored the muscle pathology of the FRG1 mouse during its life
from three weeks (no sign of disease) to 14 weeks of age (full development of the dystrophic
phenotype). We found that the onset of the phenotype is at four weeks of age when FRG1 mice display
reduced myofiber size. Muscle stem cell ex-vivo and in-vivo experiments and muscle regeneration
assays indicated that myogenic stem/progenitor cells from FRG1 mice exhibit defects in activation,
proliferative, clonogenic, differentiation and regenerative potential, suggesting that these defects
contribute to FRG1 mouse pathology. Next, we performed a gene expression profiling that identified
networks and genes affected by FRG1 over-expression that could explain the abovementioned tissue
and muscle stem cell defects.
Journal of Cell Science
FRG1 mice display a post-natal muscle growth defect.
In order to investigate the progressive course of the disease, we sacrificed FRG1 and wild-type
(WT) mice from three weeks (no phenotype) to 14 weeks of age (clear dystrophic signs) and
cryosectioned their vastus muscles. Morphometric analysis of Gomori-trichrome stained vastus
muscles revealed that the first alteration in FRG1 mice appears at four weeks of age with a reduction in
the myofiber Cross-Sectional Area (CSA) compared to WT controls (Fig. 1A; unpaired t test: p=0.0199,
n=3). Dystrophic symptoms, like infiltration of inflammatory cells and regeneration became markedly
visible between six and eight weeks of age. Flow cytometric quantification revealed an increase in the
percentage of CD45+ cells in FRG1 muscle-cell preparations, though the total number of muscle cells
isolated was not altered (Fig. 1B; unpaired t test for seven weeks: p=0.0105, n=3 and Table S1 in
supplementary material). Regeneration was assayed by quantification of centrally-nucleated myofibers
in transversally cryosectioned vastus muscles (Fig. 1C,D; paired t test for eight weeks: p=0.0129, n=6)
and regenerating myofibers were also confirmed using immunofluorescence for developmental Myosin
heavy chain (MHCd) (see Fig. S1 in supplementary material). Other muscular dystrophy signs like
necrosis, fibrosis and fat deposition were only evident at 14-weeks of age (Fig. S1 in supplementary
As shown in Figure 1A, the myofiber CSA of WT mice rapidly increased between three and
eight weeks of age as expected. Surprisingly, the myofiber CSA of FRG1 mice remained almost the
same (Fig. 1A). To further study this feature, longitudinal cryosectioning of vastus from four-weeks old
mice and quantification of the number of myonuclei per myofiber was performed. FRG1 mice
displayed a 40% reduction in the number of myonuclei compared to control mice (Fig. 1E; unpaired t
test: p=0.0122, n=3), while the number of myofibers was not altered (Fig. 1F). Hence, FRG1 mice
suffer from a post-natal muscle growth defect.
Muscle satellite cells from FRG1 mice are defective.
Although the FRG1 transgene is under the Human Skeletal Actin (HSA) promoter and thus it
should be expressed solely in mature muscle fibers, Chen and colleagues recently reported that FRG1
over-expression also occurs in primary myoblasts, causing a proliferative defect (Chen et al., 2011).
Prompted by this observation, we considered that a defect in satellite cells, the myogenic stem cells
responsible for postnatal growth and muscle repair (Bentzinger et al. 2012), could contribute to the
Journal of Cell Science
muscle growth defect of FRG1 mice. Accordingly, we found that the FRG1 transgene was expressed at
similar levels in vastus muscle and in quiescent satellite cells, freshly isolated by flow-cytometry as
SM/C-2.6+ cells (Fukada et al., 2004) (see Fig. S2 in supplementary material). Thus, in FRG1 mice the
transgene is aberrantly over-expressed in satellite cells opening the possibility to investigate the FRG1
role in this important muscle compartment. We analyzed two-weeks old animals, in which the
proliferation of satellite cells contributes to post-natal muscle growth by increasing the number of
myonuclei (Bentzinger et al.; White et al., 2010). Immunofluorescence for the paired box gene 7
(Pax7), the most important marker of quiescent satellite cells (Seale et al., 2000), and the Ki67
proliferation marker on transverse cryosections showed that the percentage of Ki67+ proliferating cells
within the Pax7+ cell population was lower in FRG1 (see Fig. S3 in supplementary material) than WT
muscles (paired t test: p=0.0271, n=3). Hence, a satellite cell impairment is present in FRG1 mice well
before the development of any histological sign of dystrophy suggesting a role for satellite cells in the
muscle growth defect of FRG1 mice.
Next, we decided to repeat this analysis ex vivo. To exclude that our observations were a
secondary effect of muscle degeneration, we analyzed mice at four weeks of age, when no dystrophic
symptoms other than a reduced myofiber size are manifested. Immunofluorescence on muscle cells
cultured for 24 hours, revealed a reduced percentage of Ki67+ within the Pax7+ cell population in FRG1
cultures compared to WT controls (Fig. 2A,B; unpaired t test: p=0.0427, n=4). Upon isolation, one of
the first steps in satellite-cell activation and exit from quiescence is the expression of MyoD (Myogenic
Differentiation-1), which controls myogenic commitment of proliferating cells (Charge and Rudnicki,
2004). Exit from quiescence is also associated with down-regulation of the quiescence marker
Caveolin-1 (Cav1) (Gnocchi et al., 2009; Volonte et al., 2005). Therefore, we monitored the temporal
expression of Pax7, MyoD and Cav1. Interestingly, 24 hours after isolation the percentage of MyoD+
cells within the Pax7+ cell population was lower in FRG1 than WT cultures (Fig. 2C,D; unpaired t test:
p=0.0029, n=3); while, the percentage of Cav1+ quiescent cells within the Pax7+ cell population was
higher in FRG1 cultures compared to WT controls (Fig. 2E,F; unpaired t test: p=0.0003, n=4). Notably,
the percentage of Pax7+ cells in FRG1 cell cultures was not decreased compared to WT (data not
shown) and we found no evidence for increased senescence or apoptosis in FRG1 satellite cells
compared to WT (Dimri et al., 1995; Kudryashova et al., 2012) (see Fig. S4 in supplementary material).
Overall, these data suggest that FRG1 satellite cells display an activation, and cell-cycle entry defect
associated with an impairment in quiescence exit.
To further support our findings, we decided to assay the clonogenic potential of FRG1-derived
Journal of Cell Science
myogenic stem/progenitor cells. Muscle cells were isolated, cultured in low-density and the number of
myogenic clones was counted. In accordance with our previous results, FRG1 cultures contained less
myogenic clones than WT (Fig. 3A; two-way Anova test: p=0.0001, n=10) resulting in a strikingly
reduced number of total MyoD+ cells (Fig. 3B; unpaired t test: p=0.0001, n=4), as assayed by
immunofluorescence. On the other hand, the clones emerging in FRG1 cultures contained a number of
cells similar to WT clones (Fig. 3C). Since the proliferative rate of FRG1 cells was only slightly lower
than WT cells (Fig. 3D; unpaired t test: p=0.0176, n=3), we concluded that the greatly reduced number
of total myogenic cells (Fig. 3B) is primarily caused by an activation defect.
Next, we analyzed the differentiation ability of primary myoblasts from FRG1 mice. To this
aim, we plated an equal number of muscle mononuclear cells before induction of differentiation, thus
avoiding a possible bias due to the reduced cell number in FRG1 initial cultures. Interestingly, we
found that FRG1 myoblasts displayed significantly reduced terminal differentiation compared to WT
(Fig. 3E,F; two-way Anova test: p=0.005, n=3), though the percentage of MHC+ cells was not reduced
in FRG1 cultures compared to WT (Fig. 3G). While this result suggests that a fusion defect could be
involved in the terminal differentiation defect of FRG1 myoblasts, further work is required to
determine the molecular mechanism underlying it.
To assure that the abovementioned defects are not caused by a reduction of the initial number of
myogenic cells, several independent approaches were employed to measure the number of quiescent
satellite cells in vivo. Firstly, the expression level Pax7 (Seale et al., 2000), was similar between FRG1
and WT mice (Fig. 4A). Secondly, the number of satellite cells defined anatomically as Pax7+
mononuclear cells located underneath the basal lamina of adult myofibers was equal between FRG1
and WT animals (Fig. 4B,C). Thirdly, muscle mononuclear cell preparations from FRG1 mice and WT
littermates contained a similar percentage of satellite cells identified by flow cytometry and cytospins,
as CD34+/Integrin-α7+/CD31-/CD45-/Sca1- and immunofluorescent-Pax7+ cells, respectively (Fig.
4D,E). Collectively, our results indicate that long before the development of muscular dystrophy FRG1
mice display myogenic stem-cell activation, proliferative, clonogenic and differentiation defects.
FRG1 mice exhibit impaired muscle regeneration.
Upon injury, satellite cells are activated, proliferate and fuse to repair the muscle (Charge and
Rudnicki, 2004). To evaluate the regeneration ability of satellite cells, muscle injury was induced by
cardiotoxin (CTX) injection in vastus of FRG1 and WT mice at four weeks of age. 10 days after CTX
injection, mice were sacrificed and their vasti were isolated and transversally cryosectioned.
Journal of Cell Science
Morphological analysis revealed that muscles from FRG1 mice contained extremely small centrally-
nucleated (regenerating) myofibers (CSA: 1009 µm2 ± SEM=119 μm2) when compared to WT controls
(CSA: 2178 μm2 ± SEM=69 μm2; Fig. 5A; unpaired test: p=0.0009, n=3). While developmental MHC
(MHCd) immunofluorescence staining was absent in WT muscles, as expected (Murphy et al., 2011),
MHCd+ regenerating myofibers were still evident in FRG1 muscles confirming a delay in muscle
regeneration (Fig. 5B). Four weeks after cardiotoxin injection, WT injured muscles, although not
completely regenerated, contained a significantly lower percentage of centrally-nucleated myofibers
than FRG1 injured muscles (Fig. 5C,D; unpaired t test: p=0.004, n=3). qRT-PCR for the muscle
regeneration marker MyHC-emb (myosin, heavy polypeptide 3, skeletal muscle, embryonic) confirmed
the delayed regeneration in FRG1 muscles (Fig. 5E; paired t test: p=0.0208, n=6). To further evaluate
the extent of this defect, we performed repeated injury experiments, where muscles were injected four
times with an interval time of one week. Contrary to WT animals, FRG1 mice were unable to repair the
damaged muscle and a part of the tissue was replaced by fat, as shown in vastus transversely
cryosectioned, four weeks after the last damage (Fig. 5F).
An in-vivo immunofluorescence analysis for Pax7 and Ki67 on muscle transverse cryosections
two days after the damage, revealed that the number of Pax7+/Ki67+ proliferating cells was lower in
FRG1 than WT muscles (Fig. 5G: paired t test: p=0.0096, n=3), suggesting a role for satellite cells in
the muscle regeneration impairment of FRG1 mice. Nonetheless, the FRG1 transgene is expressed in
both satellite cells and adult muscle, thus a reduced regeneration could be due to defects in any of these
compartments. To investigate if a cell-autonomous defect contributes to the muscle regeneration
impairment of FRG1 mice, we performed satellite cell transplantation assays. To this aim, we
generated WT and FRG1 EGFP+-transgenic mice. Freshly isolated satellite cells were transplanted in
CTX-treated tibialis anterior of WT animals to assess the ability of purified muscle stem cells to engraft
and proliferate upon transplantation into WT muscles. After three weeks, we sacrificed the mice and
analyzed cryosectioned muscles by immunofluorescence with an anti-EGFP antibody (Fig. 6A).
Although, the number of EGFP+ myofibers was similar in mouse muscles transplanted with WT and
FRG1 satellite cells (Fig. 6B), the CSA frequency distribution was different for the FRG1 donor-
derived myofibers compared to WT controls (Fig. 6C; Kolmogorov-Smirnov test: p<0.001, n=6) and
the FRG1 muscle stem cells gave rise to smaller myofibers compared to WT (Fig. 6D), indicating a
reduced ability of FRG1 satellite cells to contribute to muscle upon transplantation.
Although we cannot completely rule out the possibility of defects in other cell types or a direct
effect on the myofiber itself as potential contributing factors, these results suggest that pre-dystrophic
Journal of Cell Science
FRG1 mice display a cell-autonomous defect in the muscle stem cell compartment that could contribute
to the development of muscular dystrophy.
Expression profiling identifies pathways relevant for the pathology.
To explore the molecular mechanism underlying the disease onset, we performed a gene
expression profiling of vastus muscles from FRG1 and WT mice at four-weeks of age (n=3), when no
dystrophic symptoms other than a reduced myofiber size are present, using Illumina BeadChip
expression arrays. 390 genes were differentially expressed (up-regulated: 310; down-regulated: 80)
between FRG1 and WT samples (Fig. 7A; p<0.01; fold-difference=2). The microarrays results were
validated by qRT-PCR, using a selection of differentially expressed genes (see Table S2 in
supplementary material). Although our analysis was conducted on animals that did not present any sign
of muscular dystrophy at histological level (Fig. 1), an Ingenuity Pathway Core Analysis (IPA) of all
differentially expressed genes recognized an over-representation in several gene categories and
networks associated with muscular dystrophy, like muscle disorders, cell death, muscle development
and function and gene expression (see Table S3 in supplementary material). Intriguingly, performing a
gene-set enrichment analysis using FSHD muscle datasets available in the Gene Expression Omnibus
(GEO) database we found that FRG1 mice show a statistically significant, concordant expression
profile with FSHD patients (Fig. 7B). Furthermore, to examine the lists of genes differentially
expressed in these earlier studies, as a whole, we used rotation gene set testing (ROAST). We
investigated if these genes display a higher up-regulation or down-regulation than average universal
gene expression differences in our dataset. These tests were performed separately for each of the FSHD
muscle datasets and significant correlation for both up-regulated and down-regulated genes between
our study and the previous studies was observed (see Table S4 in supplementary material). Hence, our
results further validate the FRG1 mouse as a useful animal model of FSHD and identify pathways
relevant for its phenotype.
Journal of Cell Science
Despite its extensive study, FSHD pathogenesis remains unclear and controversial. All current
models predict that deletion of D4Z4 repeats results in the de-regulation of a candidate gene(s), located
in the FSHD region, leading to disease (Cabianca and Gabellini, 2010; van der Maarel et al., 2011).
While the two most accepted FSHD candidate genes are DUX4 and FRG1, the molecular and cellular
mechanism following their de-regulation and finally causing the disease remains elusive. Furthermore,
FSHD is characterized by an extreme variability in disease onset, progression and severity. This
heterogeneity in disease manifestation could reflect heterogeneity in gene expression of FSHD
candidate gene(s). An interesting possibility, therefore, is that the complexity of FSHD could be
explained envisaging that the epigenetic alteration of DUX4, FRG1 and other potential genes could
collaborate to determine the final phenotype.
The FRG1 mouse is a useful model of FSHD.
D4Z4 is a primate-specific repeat (Clark et al., 1996), consequently genetic mouse models of
FSHD (displaying for example a different number of subtelomeric D4Z4 repeats) cannot be generated.
Therefore, the functional consequence of D4Z4 deletion, like over-expression of an FSHD candidate
gene, is the only disease aspect that can be modeled in mice. FRG1 over-expression in mouse, frogs
and worms causes an FSHD-like phenotype (Gabellini et al., 2006; Hanel et al., 2009; Liu et al., 2010;
Wuebbles et al., 2009). Importantly, we also found a remarkable similarity of the expression profile of
FRG1 mice to the one of FSHD patients. Hence, the FRG1 mouse is to date the only mouse model that
displays features of the human disease.
Stem-cell defects could significantly contribute to the pathology of FRG1 mice.
The detailed morphological analysis of vastus muscles from FRG1 mice demonstrated that they
suffer from a postnatal muscle growth impairment, likely caused by defects of the satellite cells. In
particular, we found that long before the development of any dystrophic sign FRG1 mice display
myogenic stem cells activation, proliferative, differentiation and clonogenic defects. Satellite cells are
necessary both for the post-natal muscle growth and regeneration and indeed, cardiotoxin injection
revealed that muscle regeneration is also impaired, strengthening the importance of the
Journal of Cell Science
In the classical type of muscular dystrophies, the lack of a functional dystrophin–glycoprotein
complex, causes mechanical fragility, contraction-induced damage and death of the muscle fibers. This
leads to activation of the myogenic stem cells that take part in muscle regeneration. As a result,
dystrophic muscles undergo repeated cycles of muscle fiber degeneration and regeneration. These
vicious cycles continue until, in the late stages of the disease, the endogenous stem cell pool becomes
exhausted and muscle fibers are replaced by fibrotic and adipose tissues, compromising normal muscle
function (Mann et al., 2011). Hence, in classical muscular dystrophies, stem cells are not a primary
target of the disease; but stem-cell defects are an indirect effect of the primary mutation, which causes
muscular dystrophy, and they only contribute to disease progression.
Similarly, a proliferative defect of FRG1 primary myoblasts isolated from 18-weeks old mice
has been recently reported (Chen et al., 2011). FRG1 mice at 18 weeks of age show a pronounced
dystrophic phenotype, therefore the myoblast defect described at this age could be secondary to muscle
wasting and simply caused by the exhaustion of the muscle-stem cell proliferative capacity in the
severely dystrophic muscle. On the contrary, all our myogenic-cell and muscle regeneration
experiments were performed in young animals, well before the appearance of dystrophic symptoms.
This feature distinguishes our study, since we can exclude the possibility that the observed deficits are
secondary effects of the dystrophic phenotype, strongly suggesting that stem-cell defects can be a
primary component of muscular dystrophy in FRG1 mice.
Our data show that the FRG1 transgene is already expressed in satellite cells and in conjunction
with our transplantation experiments in WT recipient muscles, where the donor myofibers arising from
FRG1 satellite cells were smaller than the WT-derived ones, indicate a novel role for FRG1 in the
regulation of stem/progenitor cell function. Nevertheless, FRG1 satellite cells gave rise to a similar
number of donor myofibers as WT cells, suggesting that a WT myofiber and/or satellite-cell
compartment can attenuate the FRG1 muscle-stem cell defects. Thus, we do not exclude the possibility
that a mature myofiber component can play an important role in the muscle abnormalities of FRG1
mice. Indeed, the muscle microenvironment in mice can influence the behavior/abilities of myogenic
cells (Carlson and Conboy, 2007; Carlson et al., 2008; Conboy et al., 2003).
Potential molecular mechanisms for FRG1 satellite cell defects.
Our comparative gene expression profiling and the Ingenuity Pathway Core Analysis (IPA) of
all differentially expressed genes recognized a high level of over-representation in several gene
categories associated with muscular dystrophy, like inflammation, cell death, muscular disorders and
Journal of Cell Science
genetic disorders. In addition, we have identified several genes that could explain the emerging tissue
and satellite-cell defects. Interestingly, Nitric oxide synthase 1 (Nos1) was found to be down-regulated
in FRG1 muscles. Nos1 activity inhibition by pharmacological treatment or a knock-out approach
reduces the activation of satellite cells and leads to muscle regeneration deficit (Anderson, 2000;
Tatsumi et al., 2002; Tatsumi et al., 2006). Therefore, it is tempting to speculate that the FRG1 satellite
cell defects could be associated with this pathway.
Overall, this study provides a significant insight in the mechanisms that contribute to the
pathophysiology of muscular dystrophy in FRG1 mice and suggests that muscle-stem cells defects
could contribute to the disease.
Journal of Cell Science
Materials and Methods
Mouse handling, muscle injury and satellite-cell transplantations
FRG1 mice (Gabellini et al., 2006), control C57BL/6J littermates and C57BL/6-Tg
(ACTbEGFP)1Osb/J mice (a gift from Dr. Giuliana Ferrari) were maintained at Charles River (Calco,
Italy). To generate EGFP transgenic mice that over-express FRG1, C57BL/6-Tg(ACTbEGFP)1Osb/J
heterozygotes were bred with FRG1-high mice. Mice at 3–14 weeks of age were sacrificed for this
study. To induce muscle injury, 30 µl of CTX ( 0.1 mM) (Sigma) were injected into vastus lateralis
muscles of four weeks-old males using a 29G syringe. For repeated injury experiments, muscles were
injected four times with an interval time of one week, and analyzed four weeks after the last damage.
For transplantation experiments, WT males at 10–11 weeks of age were used as recipients and were
injured 48h before transplantation into tibialis anterior muscles. Muscle SM/C-2.6+/GFP+ cells were
sorted by flow cytometry as described below and 5000 cells were transplanted in each injured muscle.
Mice were sacrificed 10 days, three or four weeks after injury.
Primary muscle cell cultures, flow cytometry and sorting
Cell preparations were obtained by vastus lateralis muscles of four weeks-old males as
previously described (Xynos et al., 2011) and were plated on collagen-coated dishes after pre-plating
for 1 hour in uncoated dishes. Primary myoblasts were grown in nutrient mixture F-10 Ham (Sigma)
supplemented with 20% FBS (Hyclone) and 5ng/ml bFGF (Peprotech) for 1–5 days and differentiated
in Dulbecco’s modified Eagle medium (DMEM; EuroClone) supplemented with 5% donor horse serum
(EuroClone). To calculate the fusion index, after the initial expansion, cells were trypsinized and re-
plated in equal number before induction of differentiation, thus avoiding a possible bias due to the
reduced cell number in FRG1 initial cultures. Next, cells differentiated for 1–2 days and were stained
with anti-MHC antibodies and Hoechst. The fusion index was calculated by counting the total number
of nuclei (Hoechst stain) and the number of nuclei that are present inside a mature myotube (defined as
a MHC-positive syncytium with 3 or more myonuclei), and expressed as percentage of myonuclei. For
clonogenic assays, muscle cells were plated at 200 cells/cm2 and let grow (for 4–5 days) until visible,
well-isolated colonies were formed. Next, the total number of clones and the number of cells per clone
in the plate were counted. For proliferative assays, following an initial expansion of three days, cells
were trypsinized, re-plated in equal numbers and counted after 14, 24, 38 and 48 hours. For cytospins,
freshly isolated mononuclear cells were spotted in glass slides (20000 cells/slide) and centrifuged for 5
Journal of Cell Science
min at 800 rpm. Cells were fixed in 4% paraformaldehyde (Electron Microscopy Science) for 15 min at
room temperature and stained as described below. For flow cytometry, mononuclear cells freshly
isolated from muscles were stained with antibodies for 45 minutes at 4ºC and resuspended in DMEM
supplemented with 10% FBS (EuroClone) at a density of 10 x 106 cells/ml. The following antibodies
were used: CD45-FITC (#553080), Sca1-FITC (#553335), CD31-FITC (#553335), CD34-
AlexaFluor647 (#560233) (BD PharMingen), Integrin α7-PE (#K0046-5, MBL) and SM/C-2.6-biotin
(dilution 1/200) (Fukada et al., 2004). Flow cytometric analysis and sorting of muscle SM/C-2.6+ cells
were performed with FACSCanto (Beckton Dickinson) and MoFlo Cell Sorter (Beckman Coulter)
equipped with Argon Laser 488nm and He-Ne Laser 635nm, respectively
Muscle histology and immunofluorescence
Vastus lateralis and tibialis anterior muscles were dissected, frozen in isopentane cooled in
liquid nitrogen and cryosectioned (8-µm thick). Immunofluorescence and Gomori-trichrome and X-gal
staining were performed as previously described (Dubowitz, 1985; Xynos et al., 2011).
Characterization of fibrotic and fat tissue in the muscle was performed after Sirius Red and Oil Red O
staining, respectively (Bortolanza et al., 2011). The following primary antibodies were used: mouse
anti-MyoD1 clone 5.8A (#M3512, Dako; dilution: 1/50), rabbit anti-MyoD clone M-318 AC (sc-
760AC, SantaCruz; dilution: 1/50), rabbit anti-Cav1 (sc-894, SantaCruz; dilution: 1/50), rabbit anti-
Ki67 (NCL-Ki67p, Novocastra Laboratories; dilution: 1/200), mouse anti-MHCd (NCL-MHCd,
Novocastra; dilution: 1/40), mouse MF20 antibody (Developmental Studies Hybridoma Bank; dilution:
½), rabbit anti-GFP (A11122, Molecular Probes; dilution: 1/200), chicken anti-Laminin (ab14055,
Abcam; dilution: 1/2000), rabbit anti-Laminin (L9393, Sigma; dilution 1/300), rabbit anti-Casp3-
activated (LS-C12476, LCBio; dilution: 1/100) and mouse anti-Pax7 (hybridoma bank, dilution: ½).
Alexa Fluor 488 goat anti-rabbit, Alexa Fluor 488 goat anti-mouse, Alexa Fluor 555 goat anti-rabbit,
Alexa Fluor 555 goat anti-mouse and Alexa Fluor 555 goat anti-chicken (Molecular Probes, 1/500)
were used for secondary detection. Samples were mounted in aqueous medium and visualized at room
temperature, using Imager.M2 (N-Achroplan 10x/0.25 NA and 20x/0.45 NA) and Observer.Z1 (N-
Achroplan 10x/0.25 NA Ph1 and 20x/0.4 NA Ph2 Korr) microscopes (Zeiss). Pictures were acquired
with AxionCamMRc5 and AxioCam MRm cameras respectively using its AxioVision Rel. 4.8.2
software by Nikon. All images were analyzed with ImageJ. Adobe Photoshop C5 was used to compose
the final pictures.
Journal of Cell Science
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DNA microarray and real-time PCR analysis
Total RNA from tissues was extracted and treated with DNase 1, using the RNeasy Fibrous
Tissue Midi or Mini Kit (Qiagen). Aliquots of RNA (500 ng) samples were checked for integrity
quality number (RIN) above 8 using Agilent 2100 Byoanalyzer and amplified according to the
specifications of the Illumina TotalPrep RNA Amplification Kit (Ambion, Austin, TX, USA). The
cRNA samples were applied to the arrays of Sentrix MouseWG-6_V2 BeadChip (Illumina, San Diego,
CA, USA) and hybridized according to the manufacture’s specifications. The Sentrix BeadChips were
scanned with the Illumina's Beadarray system 500G Scanner (Illumina). The hybridization-image
signal intensity has been extracted, background subtracted and normalized using Illumina Inc.
BeadStudio software version 3.3.7. The produced data has been checked to respect the Illumina internal
quality control and loaded into Bioconductor software. All the raw data are available under the GEO
record GSE28575. cDNA was synthesized using Invitrogen’s SuperScript III First-Strand Synthesis
Super-Mix. qPCRs (for primers see Table S5 in supplementary material) were performed with SYBR
GreenER qPCR SuperMix Universal (Invitrogen) using Biorad’s CFX96 Real-time System. Relative
quantification was calculated with CFX Manager Software V.1.6. Validation of the differential
expression of genes identified by DNA microarray was performed using TaqMan gene expression
assays with custom-made TaqMan array microfluidic cards (Applied Biosystems). Relative
quantification was calculated with qBasePLUS V.1.5 using Gapdh, Ppia and 18S rRNA as reference
Statistical and bioinformatic analysis
All the microarrays raw data has been background subtracted, log2 transformed and Quantile
normalized using Bioconductor software package Lumi (Du et al., 2008). To identify differentially
expressed genes based on moderate t-test, the bioconductor Limma package (Smythe, 2005) has been
used. Genes have been selected using a p-value cut-off (after BH11 adjustment) set to the minimal
widely used p<0.01 (Shi et al., 2006) to control the false discovery rate and a log2 fold-difference
detection limit of 1. To test the association of selected differentially expressed genes with Signaling
Pathways, Cellular Process and Molecular Network, information provided in the Ingenuity Pathway
Knowledge Base has been used with a Score cut off of 3.3 (i.e p value<0.0005). Comparison with
public data has been performed using Parametric Gene Set Enrichment Analysis (PGSEA)
(Subramanian et al., 2005) and the derived GAGE and PAGE (Luo et al., 2009) and the Roast function
from Bioconductor packages. All the Public data used has been downloaded from Gene Expression
Journal of Cell Science