Duchenne muscular dystrophy (DMD) is one of the
commonest hereditary muscular diseases and is characterized
by progressivemuscle degeneration in 1 out of 3500 male live
births. The gene responsible for DMD is located on the X-
chromosome and was identified by positional cloning (Koenig
et al., 1987; Hoffman et al., 1987). It is the largest and among
the most complex of known genes, comprising 79 exons
spanning more than 2.4 million base pairs (Koenig et al., 1987).
The gene encodes at least 17 protein products of which the
main muscle isoform is among the largest, consisting of a 3685
amino acid (427 kDa) single chain that can be divided into the
following four domains; actin-binding N-terminal, rod,
cysteine-rich and C-terminal (Bies et al., 1992; Nishio et al.,
1994; Muntoni and Strong, 1989; Nudel et al., 1989; Monaco
et al., 1986; Gorecki et al., 1992; Holder et al., 1996; Feener
et al., 1989; Lidov et al., 1995; Byers et al., 1993; Austin et
al., 1995). Dystrophin protein plays a central role in organizing
a multiprotein complex at the sarcolemma, and linking
cytoskeleton proteins to extracellular matrix proteins (Monaco
et al., 1986; Holt et al., 1998; Rando, 2001). Structural and
functional analyses have assigned most of the known functions
of the protein to the two terminals and the cysteine-rich
domain, whereas the rod domain, consisting of 24 repeated
units and spanning about half the length of the protein, appears
not to be essential to function. Most DMD mutations, be they
deletions, duplications or point mutations, occur within the rod
domain, but disrupt the reading frame and therefore prevent
translation of the crucial C-terminal domain. This distinguishes
DMD from the milder Becker muscular dystrophy (BMD), in
which mutations occur in the same regions, but the mutated
transcripts retain the reading frame, and are translated into
truncated partially functional dystrophin. The mdx mouse, a
homolog of human DMD (Bulfield et al., 1984), contains a
nonsense point mutation in exon 23 (Sicinski et al., 1989),
resulting in the lack of dystrophin expression, and cycles of
muscle degeneration and regeneration.
An intriguing feature in muscles both of DMD patients and
Duchenne muscular dystrophy and the mdx mouse
myopathies reflect a lack of dystrophin in muscles.
However, both contain sporadic clusters of revertant fibers
(RFs) that express dystrophin. RF clusters expand in size
with age in mdx mice. To test the hypothesis that the
expansion of clusters is achieved through the process of
muscle degeneration and regeneration, we analyzed
muscles of mdx
mice in which degeneration and
regeneration were inhibited by the expression of micro-
dystrophins or utrophin transgenes. Postnatal RF
expansion was diminished in direct correlation to the
protective effect of the transgene expression. Similarly,
expansion of RFs was inhibited when muscle regeneration
was blocked by irradiation. However, in irradiated muscles,
irradiation-tolerant quiescent muscle precursor cells
reactivated by notexin effectively restored RF expansion.
Our observations demonstrate that revertant events occur
initially within a subset of muscle precursor cells. The
proliferation of these cells, as part of the regeneration
process, leads to the expansion of RF clusters within
degenerating muscles. This expansion of revertant clusters
depicts the cumulative history of regeneration, thus
providing a useful index for functional evaluation of
therapies that counteract muscle degeneration.
Key words: Dystrophin, Revertant fibers, Duchenne, Muscle
Expansion of revertant fibers in dystrophic mdx
muscles reflects activity of muscle precursor cells and
serves as an index of muscle regeneration
Toshifumi Yokota1,2, Qi-Long Lu3,*, Jennifer E. Morgan4, Kay E. Davies5, Rosie Fisher5, Shin’ichi Takeda6
and Terence A. Partridge1
1Muscle Cell Biology Group, Medical Research Council Clinical Science Centre, Hammersmith Hospital Campus, Imperial College School of
Medicine, London University, Du Cane Road, London, W12 0NN, UK
2Center for Genetic Medicine Research, Children’s Research Institute, Children’s National Medical Center, 111 Michigan Ave, NW, Washington, DC
3McColl Lockwood Laboratory for Muscular Dystrophy Research,Neuromuscular/ALS Center, Carolinas Medical Center, 1000 Blythe Blvd,
Charlotte, NC 28231, USA
4Department of Paediatrics, Imperial College London, The Dubowitz Neuromuscular Centre, Hammersmith Hospital, Du Cane Road, London,
W12 ONN, UK
5Department of Human Anatomy and Genetics, University of Oxford, South Parks Road, Oxford, OX1 3QX, UK
6Department of Molecular Therapy, National Institute of Neuroscience, National Center of Neurology and Psychiatry (NCNP), Ogawa-Higashi 4-1-
1, Kodaira, Tokyo 187-8502, Japan
*Author for correspondence (e-mail: email@example.com)
Accepted 28 March 2006
Journal of Cell Science 119, 2679-2687 Published by The Company of Biologists 2006
Journal of Cell Science
JCS ePress online publication date 6 June 2006
mdx mice is the presence of sporadic dystrophin-positive
muscle fibers, so called ‘revertant fibers’ (RFs) in an otherwise
dystrophin-negative background (Hoffman et al., 1990;
Nicholson, 1993; Sherratt et al., 1993). Our previous studies
showed that revertant dystrophins in muscles of mdx mice
involves massive loss of up to 30 exons, sometimes from two
non-contiguous regions. Deletions of this size are rare in DMD
or BMD, and double deletions are not reported. This is more
consistent with aberrant splicing than genomic deletion as the
mechanism for the restoration of dystrophin in RFs. This view
is bolstered by our failure to detect deletions in myonuclei of
RFs from the genomic regions coding for the missing part of
We also showed that RFs commonly form clonal clusters
that increase in size from a mean of 1 short segment at birth to
up to 100 fibers of more than 1 mm in length by 18 months
of age. Our working hypothesis is that revertant events are
initiated within a subset of muscle precursor cells that
proliferate in response to muscle degeneration and participate
in regeneration of new fibers. Expansion in the size and number
of RFs is attributable mainly to their relative resilience to
subsequent degeneration and would not be expected to be
reflected in any parallel expansion of the revertant precursors,
since these do not express dystrophin prior to fusion into fibers
and can enjoy no selective advantage from their prospects of
doing so. The low initial frequency of revertant events (around
0.1–0.01% as estimated at birth), together with this lack of
selective advantage for revertant myogenic precursors in vivo
or in tissue culture, is probably a major reason for the failure
of our attempts, like those of others (Gussoni and Kunkel,
personal communication), to establish clonal cultures of the
myoblast precursors of RFs.
As an alternative, we have explored the relationship between
expansion of RF clusters and myofiber degeneration and
regeneration in vivo using several animal models in which
degeneration or regeneration was experimentally modified.
First, we examined three strains of ‘micro’-dystrophin-
transgenic mdx mice – CS1, AX11 and M3 – generated by
Sakamoto and colleagues to test the functionality of truncated
(i.e. micro) dystrophins (Sakamoto et al., 2002; Yuasa et al.,
1998). Expression of the transgenes protects the dystrophic
muscles from degeneration with varying degrees of efficiency.
The largest transgene, CS1, almost completely prevents muscle
degeneration, whereas the smallest transgene, M3, has limited
impact. In each case, we used an antibody against the missing
rod region of dystrophin to identify ‘revertant’ muscle fibers.
We carried out similar studies of RFs in transgenic mdx mice
over-expressing full-length and mini-utrophin, both of which
have been shown to reduce dystrophic pathology markedly in
limb and diaphragm muscle (Tinsley et al., 1996; Tinsley et al.,
1998). Thus, these transgenics provide models that can be used
to evaluate the effect of muscle degeneration and regeneration
on the development and expansion of RFs. To eliminate
regeneration specifically in degenerating mdx muscles, we
applied local high-dose radiation, previously shown to block
regeneration (Wakeford et al., 1991), but not degeneration in
the mdx mouse (Pagel and Partridge, 1999), and compared the
pattern of RF expansions in irradiated versus non-irradiated
The results showed clearly that expansion of RF clusters is
dependent on muscle regeneration. The fact that reactivation of
Journal of Cell Science 119 (13)
radiation-tolerant muscle precursor cells produce newly
regenerated RFs suggests that revertant events occur initially
within a minority of muscle precursor cells. The proliferation
of these cells, as part of the regeneration process, leads to the
expansion of RF clusters within degenerating muscles. The
expansion of revertant clusters depicts the cumulative history
of regeneration, thus providing a useful index for functional
evaluation of therapies that counteract muscle degeneration.
RFs in muscles of micro-dystrophin-transgenic mice
To investigate the effects of truncated dystrophin expression on
the expansion of RF clusters, we examined micro-dystrophin-
transgenic mice CS1, AX11 and M3 (Sakamoto et al., 2002),
which contain four rod repeats (4.9 kb), three rod repeats (4.4
kb) and one rod repeat (3.9 kb), respectively (Fig. 1). Two
antibodies, P7 and MANDRA1, were used to distinguish the
revertant dystrophin from transgenic micro-dystrophin (Ellis et
al., 1990). P7, a polyclonal antibody against epitopes encoded
by exon 57 of the dystrophin gene, which is absent from the
micro-dystrophin constructs, stains only RFs (Fig. 2).
MANDRA1 is a monoclonal antibody against a C-terminal
epitope present both in RFs and in micro-dystrophins, and
therefore detects both (Fig. 2). All muscle fibers in the three
transgenic mice were stained with MANDRA1 (data not
shown). Only a small number of isolated fibers were stained
with P7, and the pattern of distribution is consistent with that
observed in muscles of young non-transgenic mdx mice. The
P7-positive fibers were therefore counted as RFs.
To understand the special distribution of both the transgenic
micro-dystrophins and revertant dystrophin, serial sections
were immunostained with both P7 and MANDRA1 antibodies.
All fibers that stained positively with P7 were also stained with
MANDRA1. Indeed, with MANDRA1, the staining intensity
in RFs was clearly stronger than in surrounding fibers
expressing only micro-dystrophin (Fig. 2). This indicates that
the RFs express both micro-dystrophin and revertant
dystrophin, and that expression of micro-dystrophin does not
suppress the expression of revertant dystrophin. Because no
antibody specific to the micro-dystrophins is available, we
cannot determine whether the levels of the transgenic
dystrophins were lower in the revertant dystrophin-positive
segments than in non-RFs.
We then compared the expression and clustering levels of
RFs in the three transgenic mice at 5 and 10 weeks of age. In
control mdx muscles, the total number of RFs and clusters both
increased markedly between 5 and 10 weeks of age, a period
of conspicuous muscle pathology (Fig. 2, Fig. 3A-C). By
comparison, both the number of RFs and clusters were
significantly fewer in AX11 and CS1 transgenic muscles at 5
weeks of age, and especially at 10 weeks of age (Fig. 2, Fig.
3A-C). The difference was most marked between control mdx
mice and CS1 mice, which are transgenic for the longest
isoform of the micro-dystrophin. Fewer RFs were detected in
muscles of M3 when compared with muscles from control mdx,
but the difference was not significant (Figs 2, 3). Interestingly,
both the total number and the maximal number of RFs per
cluster decreased with increasing age in both AX11 and CS1
transgenics (Fig. 3). Since CS1 transgenics experience no
obvious muscle degeneration during this period, it is likely that
decreased expression rather than loss of RFs is responsible,
Journal of Cell Science
Clustering of dystrophin revertant fibers
suggesting that events leading to the expression of revertant
dystrophins are reversible in some fibers.
Expansion of RFs requires muscle regeneration
To test the hypothesis that muscle regeneration plays an
important role in the process of RF expansion, we directly
compared the numbers of RFs and clusters to the percentage
of central nucleated fibers (CNF), a useful indicator of muscle
regeneration before it reaches plateau (about 80% or higher
CNF) in dystrophic muscles. The percentage of CNFs in CS1,
AX11 and M3 transgenic mice, similar to that reported by
Sakamoto et al. (Sakamoto et al., 2002), varied from 1.1%, to
23.7% to 53.4%, respectively, and was significantly lower than
the 70% observed in mdx mice at 10 weeks of age (Fig. 4A).
The increase in regeneration correlated well with the increase
in numbers of RFs and clusters from CS1 to M3 transgenics
and to mdx mice (Fig. 4B,C). This result further supports the
idea that muscle regeneration is an essential process for the
expansion of RFs.
RFs in muscles of full-length and mini-utrophin-
transgenic mdx mice
To determine whether the expression of micro-dystrophins
specifically contributes to the suppression of revertant
dystrophin in the transgenics, we examined RFs in both full-
length and mini-utrophin-transgenic mdx mice in comparison
with control mdx mice (Fig. 5). Although both transgenes
diminish muscle pathology, the expression of full-length
utrophin significantly reduced the number of centrally
nucleated regenerating fibers in tibialis anterior (TA),
extensor digitorum longus (EDL) and diaphragm muscles,
especially in young animals (Tinsley et al., 1996; Tinsley et
al., 1998). As shown in Fig. 5, the total number of RFs and
of clusters was, as expected, significantly lower in the TA
muscles of the utrophin-transgenic mice, most notably in the
mice transgenic for full-length utrophin, than the numbers in
control mdx muscles in all age groups from 1 month, to 6
months to >12 months of age. The size of RF cluster, as seen
in micro-dystrophin-transgenic mice, was also greatly
reduced in the utrophin-transgenic mice, with nearly all RFs
appearing as isolated single fibers in mice transgenic for full-
length utrophin, even in the 1-year-old animals (Fig. 5A).
However, small clusters with more than five RFs were still
frequently detected in the muscles of the same-aged mini-
utrophin-transgenic mice. The degree of expansion of RFs in
the two transgenics is therefore consistent with our previous
observations that full-length utrophin expression significantly
diminishes muscle degeneration and regeneration, and
maintains the muscles of the transgenics at nearly full
function despite the presence of some CNFs (not significantly
different from the normal control C57Bl10 mice by the age
of 5 weeks), whereas the mini-utrophin protects muscles
much less effectively (with about 10% of CNFs by the age of
5 weeks) (Tinsley et al., 1996; Tinsley et al., 1998). This
result thus suggests that it is the inhibition of muscle
degeneration and regeneration per se rather than expression
of a specific transgene that suppresses the expansion of RFs.
Fig. 1. Micro-dystrophin transgene structures in CS1, AX11 and M3
transgenics. The schematic view of micro-dystrophin transgenes
shows that CS1 (4.9 kb), AX11 (4.4 kb) and M3 (3.9 kb) encode 4, 3
and 1 rod domains, and 3, 2 and 2 hinge domains, respectively. The
size of the normal muscle dystrophin transcript is 14 kb and contains
24 rod domains and 4 hinges.
Fig. 2. Immunohistochemistry analysis of dystrophin
expression in muscles of micro-dystrophin transgenics.
Immunohistochemistry of RFs in muscles of M3, AX11
and CS1 micro-dystrophin-transgenic mice, and the mdx
mouse, with P7 antibody and MANDRA1 antibody at 5
weeks (5W) and 10 weeks (10W) of age. The P7
antibody recognizes RFs only, whereas MANDRA1
recognizes revertants and micro-dystrophin (top). Bar,
Journal of Cell Science
Expansion of RFs in mdx muscles is suppressed by
The importance of regeneration for the expansion of RFs was
further supported by the examination of mdx muscles after
irradiation. TA muscles of 3-week-old mdx mice were
irradiated with 4, 10 or 18 Gy and examined at 1 month, 3
months and 1 year afterwards (Fig. 6). In agreement with our
previous findings (Wakeford et al., 1991; Pagel and Partridge,
1999; Morgan et al., 2002), irradiation suppressed fiber
regeneration in a dose-dependent manner, and 18 Gy
irradiation abolished nearly all regeneration activity as
demonstrated by the lack of fibers of small caliber and by the
lack of central nucleation one month after irradiation (Fig. 6A).
The number of RFs was inversely correlated with the dosage
of irradiation. After irradiation with 18 Gy, nearly all RFs were
singular and the number of RFs in the muscles at 1 and 3
months after the irradiation remained indistinguishable from
those seen in the muscles at the time of irradiation (3 weeks)
(Fig. 6B). The suppressive effect of irradiation on RF
proliferation was most evident 1 year after irradiation (Fig.
6C). This reflected the continued growth in the number of RFs
and the size of RF clusters in non-irradiated mdx muscles,
whereas in the irradiated muscles such numbers remained at
similar levels to those of mice at 3 weeks of age. These results
provide compelling evidence that the expansion of RFs relies
on muscle regeneration.
Reactivation of muscle regeneration by notexin
treatment is associated with formation of new RFs
Our previous experiments have shown that most muscle
precursor cells are radiation sensitive and that muscle
regeneration in the mdx mouse is blocked by irradiation with
18 Gy. However, some muscle precursor cells, perhaps
quiescent satellite cells, in irradiated muscles can be
reactivated by severe injury mediated by treatment with
notexin, a snake venom, and contribute to muscle regeneration
(Heslop et al., 2000). We therefore investigated the association
between activation of such precursor cells and generation of
RFs and revertant clusters. The legs of mdx mice were
irradiated with 18 Gy at 3 weeks of age followed by notexin
injection into TA muscles 4 weeks later (Fig. 7A). As expected,
the muscles, examined 1 week after injection, showed foci of
regenerating fibers of small caliber in the notexin-treated areas.
The total number of RFs in notexin-treated and pre-irradiation
muscles was lower than that in the non-irradiated control
muscles, but significantly higher than that in the irradiated
Journal of Cell Science 119 (13)
Fig. 3. The number of RFs and clusters are decreased in
micro-dystrophin-transgenic mice. (A) The maximal number
of RFs. (B) The maximal number of revertant clusters
containing more than one RF. (C) The maximal number of
RFs in one cluster. Values are mean ± s.e.m. for 4–10 TA
muscles per group. *Significant difference (P<0.05) from mdx
muscles of the same age. 5W, 5 weeks; 10W, 10 weeks.
% of centrally
Percentage of centrally nucleated fibres
Percentage of centrally nucleated fibresM3AX11CS1
Fig. 4. Muscle degeneration and regeneration is required
for RF expansion. (A) The percentage of central nucleated
fibers in each strain of mice. The number of regenerating
fibers decreases from mdx, to M3, to AX11 to CS1 mice
aged 5 weeks and 10 weeks. (B) Correlation between the
number of RFs and the percentage of centrally nucleated
fibers in each strain at 10 weeks of age. (C) Correlation
between the number of clusters and the percentage of
central nucleated fibers in each strain at 10 weeks of age.
The direct correlation between the number of RFs or
clusters and centrally nucleated fibers suggests that the
muscle degeneration and regeneration is an essential step
in the expansion of RFs. Values are mean ± s.e.m. for 4-
10 muscles per group. *Significant difference (P<0.05)
from mdx muscles of the same age.
Journal of Cell Science
Clustering of dystrophin revertant fibers
muscles without notexin treatment (Fig. 7B,C). Most notable
is the formation of clusters of dystrophin-positive fibers within
the regenerating areas demonstrated by immunohistochemistry
(Fig. 7B). All these fibers were small in caliber and centrally
nucleated, and therefore were newly regenerated RFs.
Although notexin treatment in non-irradiated muscle did not
increase the total number of RFs significantly, it did create
some clusters of newly regenerating RFs (Fig. 7B,C). These
results show that at least some of the radiation-resistant muscle
precursor cell populations are able to establish revertant
Revertant dystrophins, but not micro-dystrophins,
restore neuronal nitric oxide synthase expression at the
Our previous studies showed that most revertant dystrophins
are able to relocalize nitric oxide synthase (nNOS) to
sarcolemma of the fibers (Lu et al., 2000). By contrast, nNOS
was not restored to the sarcolemma of muscle fibers in mice
transgenic for micro-dystrophin (Fig. 8). We therefore
examined RFs in the muscles of these transgenic mice to see
if the expression of micro-dystrophins affects the localization
of nNOS in RFs. Serial sections from all three transgenics
showed that most RFs (positively stained with antibody P7)
were also positively stained for nNOS. This suggests that most
revertant dystrophins are functionally superior in this respect
to the micro-dystrophins, and that interaction of nNOS with
revertant dystrophins is not hampered by the presence of over-
Although it was first described 15 years ago (Hoffman et al.,
1990; Sherratt et al., 1993), the phenomenon of revertant
muscle fibers remains enigmatic. In particular, we do not have
a firm idea of the mechanism behind the expression of
dystrophin in these fibers. We have argued previously (Lu et
al., 2000) that the revertant phenomenon is an epigenetic event
that arises in individual satellite cells at around birth. Here,
from the study of the pattern of appearance and expansion of
RFs in the dystrophic mice, we derive some further support for
Muscle regeneration is essential for RF expansion
One of the most prominent features of the RFs in the mdx
mouse is their expansion with age, which appears clonal in that
the pattern of epitope loss is variable between clusters but is
conserved within each cluster (Lu et al., 2000). RFs first appear
at around birth as short segments, some 10 ?m in length, of
sporadic single fibers, expanding over the following 18 months
up to clusters with more than 100 fibers and spanning 1 mm
or more of fiber length, although never attaining a sufficient
proportion to ameliorate muscle pathology significantly. This
expansion has been modeled as a consequence of cycles of
muscle degeneration and regeneration in combination with a
preferential survival of the fibers that contain dystrophin (Lu
et al., 2000; Garcia et al., 1999). However, this presumption
has never been tested and this was a major goal of this
investigation. All data in our study point to a strong and direct
dependence of RF cluster expansion on the intensity of muscle
regeneration. In mdx
mice carrying micro-dystrophin
transgenes, the number of RFs is strongly correlated with other
measures of the protection afforded against muscle damage by
each individual transgene. Similarly, in the utrophin-transgenic
mdx mice, where muscle degeneration and consequent
regeneration is diminished, there was a clear relationship
between the protective effect of the two versions of the
transgenes and the increase in size of RF clusters. The
dependence of RF expansion on muscle regeneration was
further confirmed by the effects of irradiation on RFs. By
varying the dose of radiation and by examining the target
muscles at different time points after irradiation, we found that
expansion of RFs was diminished by radiation in a dose-
dependent manner and that, at the highest dose, RFs were, in
effect, frozen at the cluster size they would have attained at the
time of application of the radiation (Fig. 6). This was most
clearly seen 1 year after irradiation with 18 Gy, when the
disparity between irradiated and control muscles had become
more conspicuous. In combination, these data clearly establish
that muscle regeneration is an absolute requirement for
expansion of RF clusters.
Myogenic stem cells with altered splicing in the gene
encoding dystrophin are the probable sources of RF
The dependence of RF expansion on regeneration argues
strongly against the mechanism we had raised previously:
namely that the expansion represents the progressive increase
in territory of factors each of which determines a specific
Fig. 5. Immunohistochemistry analysis of dystrophin
expression in muscles of utrophin-transgenic mdx mice.
(A) Immunohistochemistry with anti-utrophin and anti-
dystrophin antibodies in mdx mice and transgenic mdx
mice expressing full-length utrophin (utro/mdx) at 1 year
of age (note that a proportion of fibers are centrally
nucleated in the muscle of mice of the same age that are
transgenic for full-length utrophin). Only a single RF is
seen in the transgenic mouse compared with a large
cluster in the control mdx mouse. (B) The number of RFs
in mdx mice (black columns), mdx mice expressing mini-
utrophin (gray columns) and mdx mice expressing full-
length utrophin (white columns), at 1 month, 6 months
and 1 year of age. Each point is the mean ± s.e.m. of 3-8
muscles per group. *Significant difference (P<0.05)
from mdx muscles of the same age. Bar, 100 ?m.
Journal of Cell Science
pattern of alternative splicing, and spreads by diffusion within
each fiber and between adjacent fibers (Lu et al., 2000). This
hypothesis would predict that revertant clusters would grow
within the existing stable muscle fibers. However, the present
findings that expansion of RFs does not occur in the absence
of regeneration, even when degeneration continues (after
irradiation), argue unequivocally
participation of a diffusible factor for the expansion of RFs
Journal of Cell Science 119 (13)
Fig. 6. Reduced expression of revertant dystrophin in irradiated mdx
muscles. (A) Mice were irradiated at the age of 3 weeks with 4 Gy,
10 Gy, or 18 Gy, and sacrificed at 3 months after the irradiation. The
irradiated muscles show reduced number of RFs compared with
untreated control mdx muscle (A), suggesting that muscle
regeneration is crucial for RF expansion. (B) The number of RF
clusters, total number of RFs, and maximal number of RFs in one
cluster in mdx TA muscle at 4 weeks after irradiation at 3 weeks of
age. The mice were irradiated with 4 Gy (gray bars), 10 Gy (dashed
bars) or 18 Gy (white bars), or were non-irradiated (black bars).
(C) Comparison of the level of suppression of RF expansion at 1
month or 1 year after 18 Gy irradiation at the age of 3 weeks. Black
bars, non-irradiated muscles; white bars, irradiated muscles. Each
point is the mean ± s.e.m. of 3-8 muscles per group. *Significant
difference (P<0.05) from mdx muscles of the same age. Bar, 100 ?m.
Fig. 7. Notexin injection after irradiation restores RF expansion.
(A) Experimental design of irradiation and notexin treatments in TA
muscles of mdx mice. (1) Irradiation only; (2) notexin treatment
only; and (3) notexin treatment 4 weeks after 18 Gy irradiation at 3
weeks of age. IHC, immunohistochemistry. (B) The irradiated TA
muscle shows a single RF, in contrast to the untreated control TA
muscle and notexin-injected muscles without irradiation (notexin
only). A regenerating RF cluster reappears after notexin injection in
pre-irradiated muscles (18 Gy + notexin). Bar, 100 ?m. (C) The total
number of RFs and the maximal number of RFs per cluster in TA
muscle after notexin treatment with or without pre-irradiation.
*Significant difference (P<0.05) was obtained between control mice
treated with 18 Gy irradiation only and the mice treated with notexin
after 18 Gy pre-irradiation. Each point is the mean ± s.e.m. of 3-4
muscles per group.
Journal of Cell Science
Clustering of dystrophin revertant fibers
within dystrophic muscles. The clonal individuality of epitope
expression (Lu et al., 2000) and the dependence on muscle
regeneration therefore indicate clearly the involvement of
myogenic stem cells for the expansion of RFs.
In principle, translatable transcripts might be generated by
either further mutations that restore an open-reading frame, or
by skipping of frame-disrupting exon(s) during splicing of the
transcript. The clonal individuality of the RF cluster favors the
mutation hypothesis (Klein et al., 1992; Wallgren-Pettersson et
al., 1993), but the high frequency of revertant events and of
clusters in which exons are lost from two non-adjacent parts of
the gene encoding dystrophin argue against this view (Lu et al.,
2000). A random mutational process would have been expected
to generate at least some large multi-fiber domains at birth,
arising from mutations occurring early during prenatal
myogenesis. We found no such clusters at birth. Furthermore,
we could not demonstrate deletion of the genomic sequence
corresponding to the missing epitopes in the protein sequence
within large RF clusters. Results from our present studies now
provide more-compelling evidence against the mechanism of
superimposed mutations (Fig. 3) (Crawford et al., 2001),
leaving an epigenetic mechanism as the most plausible
explanation (Wilton et al., 1997; Lu et al., 2000). All transgenic
mdx mice showed similar numbers of RFs before the onset of
muscle degeneration and nearly all of these RFs were singular.
Furthermore, no expansion of RFs was observed in any
transgenic mdx mouse despite a significant muscle growth
before the age of 5 weeks. Instead, the number of RFs and the
intensity of dystrophin signals decrease with age in mini-
dystrophin- and micro-dystrophin-transgenic mdx
suggesting that at least some of the revertant phenotypes are
not stably maintained and could still be subject to modulation.
Because no counterpart of such a phenomenon has been
described in other biological systems, the mechanism is still
open to speculation, but seems likely to involve alteration in
splicing. The dystrophin gene and expansion of RFs could be
a unique model for exploring mechanisms in the evolution of
In situ model for assessing muscle regeneration
Skeletal muscle has considerable regenerative capacity both in
response to injury and in the recurrent dystrophic process.
However, it is difficult to measure regenerative activity in vivo.
The clonal expansion of RFs provides a unique tool for
assessing the regenerative capacity of the muscle. Before the
onset of muscle degeneration in mdx mice, dystrophin in nearly
all RFs spans short segments of single fibers, corresponding in
size to a single nuclear domain. The fact that these domains
subsequently expand along the fiber and to neighboring fibers
is, as we show here, best explained by the presence of an
adjacent myogenic precursor harboring the revertant event.
One year later, RF clusters have grown up to 100 fibers across
and over 1 mm in length (Lu et al., 2000). From counts of
myonuclear density, we estimate that a cluster of such size
contains some thousands of myonuclei. The clonal nature of a
revertant cluster implies that most of these thousands of
myonuclei are the offspring of the single myogenic cell that
was present at birth. The presence of myogenic cells with such
mobility, and sufficient capacity to produce a large number of
fibers in adult muscles in vivo, fits well with the evidence from
our recent myofiber implantation experiments (Collins et al.,
2005). When isolated fibers were transplanted into host
muscles, we demonstrated that a single myofiber bearing on
average fewer than seven satellite cells can produce a cluster
of some 300 fibers, resembling in many respects the clusters
of RFs in mdx mice. Similarly, we have demonstrated
previously that the majority of satellite cells lose proliferative
capability after irradiation with 18 Gy, but that a tiny subset of
radiation-resistant myogenic precursors, constituting probably
in the region of 1% of the radiation-sensitive population,
retains the ability to restitute the satellite cell population
partially (Heslop et al., 2000). The appearance of RF clusters
in notexin-treated pre-irradiated muscles suggests that some of
the revertant myogenic cells belong to this radiation-resistant
precursor population. Expansion of RFs in mdx mice therefore
provides potentially useful models for studies of myogenic
Expansion of RFs serves as a cumulative index of
pathological activity in the dystrophic muscles
The link between regeneration and expansion of revertant
clusters allows us to use the degree of expansion as a
cumulative index of past pathological activity in dystrophic
muscles. At present, the most commonly used histological
indicator of disease progression in the mdx mouse is the
progressive increase in the proportion of centrally nucleated
muscle fibers. However, this index rises very rapidly after the
onset of the dystrophy and reaches a plateau of ~80% in about
6 weeks, becoming insensitive to any continuation of
Fig. 8. nNOS expression is only found in RFs in micro-dystrophin-
transgenic muscles and mdx mice. The expression of nNOS and
dystrophin RFs in M3 and AX11 transgenic mice, and in mdx mice,
aged 5 weeks. nNOS is expressed only in RFs in these transgenics,
suggesting that the dystrophin rod domain has an important role in
targeting nNOS at the sarcolemma. Bar, 50 ?m.
Journal of Cell Science
pathology beyond this point in the dystrophic mdx mouse,
which is the most widely used model for testing of new
therapies. By contrast, the increase in size of RF clusters
continues into the second year (Lu et al., 2000), confirming that
the disease remains active into the later life of this animal
model. This should, in principle, make it useful as a means of
evaluating the beneficial effects of any treatment designed to
combat muscle fiber degeneration. This view is vindicated in
the present work by the clear concordance between the effects
of the micro-dystrophin and utrophin transgenes on the overall
measures of pathological change and on the suppression of RF
cluster expansion. Thus, the degree of expansion of the
revertant clusters in the mdx mice is by far the best available
histological indicator for assessment of long-term functional
value by therapeutic transgenes or other treatments that are
designed to combat muscle degeneration. It has the additional
advantage that it might be applicable in the dystrophic dog, in
which central nucleation is not a persistent feature of the
It is also noteworthy that the majority of RFs were able to
localize nNOS to the fiber membrane, whereas none of the
micro-dystrophins nor the utrophin exhibited this function,
supporting previous findings that the structure of part of the
rod sequence is implicated in the association with nNOS (Wells
et al., 2003). Because the RF number diminished in the
background of the most effective micro-dystrophins, it would
appear that nNOS function is not crucial to myofiber survival
if other functions of dystrophin are adequately accomplished.
In summary, our studies show that initial formation of RFs
in mdx mice is independent of dystrophic phenotype and that
alternative splicing in myogenic stem cells is likely to be
responsible for the production of revertant dystrophin and
clonal expansion of RFs. The expansion of RFs is closely
related to muscle regeneration and therefore the quantity and
distribution of RFs is a useful indicator of the long-term
functionality of therapeutic interventions. The study of
revertant dystrophin and expansion of RFs in the mdx mouse
could provide a unique model to understand the mechanisms
for the regulation of alternative splicing.
Materials and Methods
This study used mdx mice, mdx mice transgenic for CS1, AX11 and M3 micro-
dystrophin transgenes, mdx mice transgenic for full-length and mini-utrophin
transgenes, and C57/BL10 and C57/BL6 mice as controls (Sakamoto et al., 2002;
Vainzof et al., 1995; Tinsley et al., 1996; Tinsley et al., 1998).
The monoclonal antibody MANDRA1 was used to detect both RFs and micro-
dystrophins (1:100). The polyclonal antibody against neuronal nitric oxide synthase
(nNOS) (Santa Cruz) was also used (1:20).
P7 rabbit polyclonal antibody was raised against a polypeptide within exon 57 of
the gene encoding dystrophin. P7 antibody detects only RFs and not the micro-
dystrophins (1:1000). The anti-utrophin polyclonal antibody (G3; 1:25) was used
for immunohistochemistry (Tinsley et al., 1998).
Acetone-fixed cryosections (6 ?m) were incubated with primary antibody at 4°C
overnight. Alexa Fluor 488- or 594-labeled goat anti-rabbit IgG (H+L) (Molecular
Probes) was used as the secondary antibody (1:200). For immunofluorescent
staining, sections after the secondary antibody were counterstained with 4?,6-
diamidino-2-phenylindole (DAPI; Sigma). The sections were viewed and
photographed using a Leica microscope with METAMORPHTMImaging system.
For immunoperoxidase staining, the primary P7 antibody was applied in antibody
diluent solution (Dako) at 4°C overnight. After washing, sections were incubated
with a biotin-conjugated anti-rabbit IgG (1:200; Dako) for 60 minutes at room
temperature, and treated with an avidin-biotin-horseradish peroxidase complex
(Dako) for 30 minutes and visualized by 3,3?-diaminobenzidine tetrahydrochloride
containing 0.015% hydrogen peroxide.
Muscle fibers were regarded as dystrophin positive only when more than half the
membrane circumference was stained in cross-sections. RFs adjacent to each other
were characterized as a single cluster. For closer comparison of RFs in mice of
different groups, serial sections (6 ?m thick) of TE muscle of every 100 ?m were
stained with antibodies. The maximal number of RFs, the number of revertant
clusters, and the number of RFs in each cluster were counted and compared.
Irradiation and Notexin treatment
mdx mice were irradiated at the age of 3 weeks. Animals were anesthetized with
subcutaneous injection of 50 ?l of hypnorm (Janssen; fentanyl citrate, final
concentration 0.79 mg/ml; fluanisone, final concentration 2.5 mg/ml) and hypnovel
(Roche; midazolam, final concentration 1.25 mg/ml). Animals were restrained
during irradiation, the legs were irradiated with 4, 10 and18 Gy delivered by an IBL
637 cell irradiator as described previously (Gross et al., 1999). Non-irradiated
contralateral legs and age-matched mdx were used as controls. At each of a series
of time points, 1, 4, 12 weeks and 1 year after irradiation, mice were euthanized
and muscles examined. For notexin treatment, mice were anesthetized as above and
the TA muscles of their right legs that had been pre-irradiated with 18 Gy 4 weeks
before were injured by injection of 10 ?l of 10 ?g/ml notexin (Heslop et al., 2000).
The injected mice were killed 7 days after the notexin injection and its TA muscles
taken for histopathology and immunohistochemistry. Three to eight samples were
examined for each treatment regime and untreated controls.
The data between samples were compared using Student’s t test. P<0.05 was
considered statistically significant.
This work was supported by grants from the Medical Research
Council, the Muscular Dystrophy Group of Great Britain, the Leopold
Muller Foundation (UK), Grant-in-Aid for Scientific Research
(KAKENHI), Grant-in-Aid for Japan Society of the Promotion of
Science (JSPS) Fellows (Japan), Aktion Benni & Co EV (Germany)
and the Neuromuscular/ALS Center (Carolinas Medical Center,
Charlotte, NC). We thank G. E. Morris (MRIC Biochemistry Group,
The North East Wales Institute, Wrexham, UK) for supplying the
monoclonal antibodies. T.Y. is a research fellow of JSPS.
Austin, R. C., Howard, P. L., D’Souza, V. N., Klamut, H. J. and Ray, P. N. (1995).
Cloning and characterization of alternatively spliced isoforms of Dp71. Hum. Mol.
Genet. 4, 1475-1483.
Bies, R. D., Phelps, S. F., Cortez, M. D., Roberts, R., Caskey, C. T. and Chamberlain,
J. S. (1992). Human and murine dystrophin mRNA transcripts are differentially
expressed during skeletal muscle, heart, and brain development. Nucleic Acid Res. 20,
Bulfield, G., Siller, W. G., Wight, P. A. and Moore, K. J. (1984). X chromosome-linked
muscular dystrophy (mdx) in the mouse. Proc. Natl. Acad. Sci. USA 81, 1189-1192.
Byers, T. J., Lidov, H. G. W. and Kunkel, L. M. (1993). An alternative dystrophin
transcript specific to peripheral nerve. Nat. Genet. 4, 77-81.
Collins, C. A., Olsen, I., Zammit, P. S., Heslop, L., Petrie, A., Partridge, T. A. and
Morgan, J. E. (2005). Stem cell function, self-renewal, and behavioral heterogeneity
of cells from the adult muscle satellite cell niche. Cell 122, 289-301.
Crawford, G. E., Lu, Q. L., Partridge, T. A. and Chamberlain, J. S. (2001).
Suppression of revertant fibers in mdx mice by expression of a functional dystrophin.
Hum. Mol. Genet. 10, 2745-2750.
Ellis, J. M., Man, N. T., Morris, G. E., Ginjaar, I. B., Moorman, A. F. and van
Ommen, G. J. (1990). Specificity of dystrophin analysis improved with monoclonal
antibodies. Lancet 336, 881-882.
Feener, C. A., Koenig, M. and Kunkel, L. M. (1989). Alternative splicing of human
dystrophin mRNA generates isoforms at the carboxy terminus. Nature 338, 509-511.
Garcia, L., Peltekian, E., Pastoret, C., Israeli, D., Armande, N. and Parrish, E. (1999).
Computerised dystrophic muscle simulator: prospecting potential therapeutic strategies
for muscle dystrophies using a virtual experimental model. J. Gene Med. 1, 43-55.
Gorecki, D. C., Monaco, A. P., Derry, J. M. J., Walker, A. P., Barnard, E. A. and
Barnard, P. J. (1992). Expression of four alternative dystrophin transcripts in brain
regions regulated by different promoters. Hum. Mol. Genet. 1, 505-510.
Gross, J. G., Bou-Gharious, G. and Morgan, J. E. (1999). Potentiation of myoblast
transplantation by host muscle irradiation is dependent on the rate of radiation delivery.
Cell Tissue Res. 298, 371-375.
Heslop, L., Morgan, J. E. and Partridge, T. A. (2000). Evidence for a myogenic stem
cell that is exhausted in dystrophic muscle. J. Cell Sci. 113, 2299-2308.
Journal of Cell Science 119 (13)
Journal of Cell Science
Clustering of dystrophin revertant fibers
Hoffman, E. P., Brown, R. H., Jr and Kunkel, L. M. (1987). Dystrophin: the protein
product of the Duchenne muscular dystrophy locus. Cell 51, 919-928.
Hoffman, E. P., Morgan, J. E., Watkins, S. C. and Partridge, T. A. (1990). Somatic
reversion/suppression of the mouse mdx phenotype in vivo. J. Neurol. Sci. 99, 9-25.
Holder, E., Maeda, M. and Bies, R. D. (1996). Expression and regulation of the
dystrophin Purkinje promoter in human skeletal muscle, heart, and brain. Hum. Genet.
Holt, K. H., Lim, L. E. and Campbell, K. P. (1998). Functional rescue of sarcoglycan
complex in the B10 14.6 hamster using d-sarcoglycan gene transfer. Mol. Cell 1, 841-
Klein, C. J., Coovert, D. D., Bulman, D. E., Ray, P. N., Mendell, J. R. and Burghes,
A. H. (1992). Somatic reversion/suppression in Duchenne muscular dystrophy (DMD):
evidence supporting a frame-restoring mechanism in rare dystrophin-positive fibers.
Am. J. Hum. Genet. 50, 950-959.
Koenig, M., Hoffman, E. P., Bertelson, C. J., Monaco, A. P., Feener, C. and Kunkel,
L. M. (1987). Complete cloning of the Duchenne muscular dystrophy (DMD) cDNA
and preliminary genomic organization of the DMD gene in normal and affected
individuals. Cell 50, 509-517.
Lidov, H. G. W., Selig, S. and Kunkel, L. M. (1995). Dp140: a novel 140 kDA CNS
transcript from the dystrophin locus. Hum. Mol. Genet. 4, 329-335.
Lu, Q. L., Morris, G. E., Wilton, S. D., Ly, T., Artem’yeva, O. V., Strong, P. and
Partridge, T. A. (2000). Massive idiosyncratic exon skipping corrects the nonsense
mutation in dystrophic mouse muscle and produces functional revertant fibers by clonal
expansion. J. Cell Biol. 148, 985-996.
Monaco, A. P., Neve, R. L., Coletti-Feener, C., Bertelson, C. J., Kurnit, D. M. and
Kunkel, L. M. (1986). Isolation of candidate cDNA clones for portions of the
Duchenne muscular dystrophy gene. Nature 323, 646-650.
Morgan, J. E., Gross, J. G., Pagel, C. N., Beauchamp, J. R., Fassati, A., Thrasher, A.
J., Di Santo, J. P., Fisher, I. B., Shiwen, X., Abraham, D. J. et al. (2002). Myogenic
cell proliferation and generation of a reversible tumorigenic phenotype are triggered
by preirradiation of the recipient site. J. Cell Biol. 157, 693-702.
Muntoni, F. and Strong, P. (1989). Transcription of the dystrophin gene in Duchenne
muscular dystrophy muscle. FEBS Lett. 252, 95-98.
Nicholson, L. V. (1993). The “rescue” of dystrophin synthesis in boys with Duchenne
muscular dystrophy. Neuromuscul. Disord. 3, 525-531.
Nishio, H., Takeshima, Y., Narita, N., Yanagawa, H., Suzuki, Y., Ishikawa, Y.,
Minami, R., Nakamura, H. and Matsuo, M. (1994). Identification of a novel first
exon in the human dystrophin gene and a new promoter located more than 500 kb
upstream of the nearest known promoter. J. Clin. Invest. 94, 1037-1042.
Nudel, U., Zuk, D., Einat, P., Zeelon, E., Levy, Z., Neuman, S. and Yaffe, D. (1989).
Duchenne muscular dystrophy gene product is not identical in muscle and brain. Nature
Pagel, C. N. and Partridge, T. A. (1999). Covert persistence of mdx mouse myopathy
is revealed by acute and chronic effects of irradiation. J. Neurol. Sci. 164, 103-116.
Rando, T. A. (2001). The dystrophin-glycoprotein complex, cellular signaling, and the
regulation of cell survival in the muscular dystrophies. Muscle Nerve 24, 1575-1594.
Sakamoto, M., Yuasa, K., Yoshimura, M., Yokota, T., Ikemoto, T., Suzuki, M.,
Dickson, G., Miyagoe-Suzuki, Y. and Takeda, S. (2002). Micro-dystrophin cDNA
ameliorates dystrophic phenotypes when introduced into mdx mice as a transgene.
Biochem. Biophys. Res. Commun. 293, 1265-1272.
Sherratt, T. G., Vulliamy, T., Dubowitz, V., Sewry, C. A. and Strong, P. N. (1993).
Exon skipping and translation in patients with frameshift deletions in the dystrophin
gene. Am. J. Hum. Genet. 53, 1007-1015.
Sicinski, P., Geng, Y., Ryder-Cook, A. S., Barnard, E. A., Darlison, M. G. and
Barnard, P. J. (1989). The molecular basis of muscular dystrophy in the mdx mouse:
a point mutation. Science 244, 1578-1580.
Tinsley, J. M., Potter, A. C., Phelps, S. R., Fisher, R., Trickett, J. I. and Davies, K.
E. (1996). Amelioration of the dystrophic phenotype of mdx mice using a truncated
utrophin transgene. Nature 384, 349-353.
Tinsley, J., Deconinck, N., Fisher, R., Kahn, D., Phelps, S., Gillis, J. M. and Davies,
K. (1998). Expression of full-length utrophin prevents muscular dystrophy in mdx
mice. Nat. Med. 4, 1441-1444.
Vainzof, M., Passos-Bueno, M. R., Man, N. and Zatz, M. (1995). Absence of correlation
between utrophin localization and quantity and the clinical severity in
Duchenne/Becker dystrophies. Am. J. Med. Genet. 58, 305-309.
Wakeford, S., Watt, D. J. and Partridge, T. A. (1991). X-irradiation improves mdx
mouse muscle as a model of myofiber loss in DMD. Muscle Nerve 14, 42-50.
Wallgren-Pettersson, C., Jasani, B., Rosser, L. G., Lazalou, L. P., Nicholson, L. V.
and Clark, A. (1993). Immunohistochemical evidence for second or somatic mutations
as the underlying cause of dystrophin expression in isolated fibers in Xp21 muscular
dystrophy of Duchenne-type severity. J. Neurol. Sci. 118, 56-63.
Wells, K. E., Torelli, S., Lu, Q., Brown, S. C., Partridge, T., Muntoni, F. and Wells,
D. J. (2003). Relocalization of neuronal nitric oxide synthase (nNOS) as a marker for
complete restoration of the dystrophin associated protein complex in skeletal muscle.
Neuromuscul. Disord. 13, 21-31.
Wilton, S. D., Dye, D. E. and Lainy, N. G. (1997). Dystrophin gene transcripts skipping
the mdx mutation. Muscle Nerve 20, 728-734.
Yuasa, K., Miyagoe, Y., Yamamoto, K., Nabeshima, Y., Dickson, G. and Takeda, S.
(1998). Effective restoration of dystrophin-associated proteins in vivo by adenovirus-
mediated transfer of truncated dystrophin cDNAs. FEBS Lett. 425, 329-336.
Journal of Cell Science