2298 The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
Assessment of disease activity in muscular
dystrophies by noninvasive imaging
Katie K. Maguire,1 Leland Lim,2 Sedona Speedy,1 and Thomas A. Rando1,2,3
1Department of Neurology and Neurological Sciences, Stanford University School of Medicine, Stanford, California, USA.
2Neurology Service and RR&D Center of Excellence, Veterans Affairs Palo Alto Health Care System, Palo Alto, California, USA.
3Glenn Laboratories for the Biology of Aging, Stanford University School of Medicine, Stanford, California, USA.
Muscular dystrophies are a class of disorders that cause progressive muscle wasting. A major hurdle for dis-
covering treatments for the muscular dystrophies is a lack of reliable assays to monitor disease progression
in animal models. We have developed a novel mouse model to assess disease activity noninvasively in mice
with muscular dystrophies. These mice express an inducible luciferase reporter gene in muscle stem cells. In
dystrophic mice, muscle stem cells activate and proliferate in response to muscle degeneration, resulting in
an increase in the level of luciferase expression, which can be monitored by noninvasive, bioluminescence
imaging. We applied this noninvasive imaging to assess disease activity in a mouse model of the human disease
limb girdle muscular dystrophy 2B (LGMD2B), caused by a mutation in the dysferlin gene. We monitored the
natural history and disease progression in these dysferlin-deficient mice up to 18 months of age and were able
to detect disease activity prior to the appearance of any overt disease manifestation by histopathological anal-
yses. Disease activity was reflected by changes in luciferase activity over time, and disease burden was reflected
by cumulative luciferase activity, which paralleled disease progression as determined by histopathological
analysis. The ability to monitor disease activity noninvasively in mouse models of muscular dystrophy will be
invaluable for the assessment of disease progression and the effectiveness of therapeutic interventions.
Muscular dystrophies are a class of inherited muscle disorders
that are characterized by progressive muscle weakness and wast-
ing. These diseases often result from mutations of genes that
are critical for muscle cell structure or function (1). Therapeu-
tic strategies to treat muscular dystrophies, including gene ther-
apies and small molecule therapies, are being investigated, but
currently there are few treatments available and none that sub-
stantially alter disease progression (2). Many dystrophic animal
models exist and provide valuable resources for understanding
the disease pathogenesis and for testing therapeutic interven-
tions (3). One of the major limitations to the study of therapeutic
agents for the treatment of muscular dystrophies is the absence of
reliable assays of disease activity in living animals.
The gold standard for monitoring disease progression or the
response to treatments in animals is the analysis of muscle histo-
pathology. This approach is labor intensive, difficult to quantify,
and usually terminal for the experimental animal. As such, inves-
tigators have sought methods to assess disease activity or progres-
sion using noninvasive or minimally invasive methods. In animal
models, levels of serum biomarkers, strength measurements, and
MRI evaluations have been used to assess disease activity and pro-
gressive deterioration of dystrophic muscle (4–6). However, these
techniques are either highly variable (especially serum biomark-
ers), nonspecific (especially strength measurements), expensive
(especially MRI), or some combination of the three. As such, there
remains a critical need for a method to provide quantitative and
reliable assessment of ongoing and cumulative disease activity
that closely reflects the histopathological changes occurring in
the muscle in dystrophic animal models.
In this report, we describe a novel mouse model in which muscle
regeneration, reflecting the response to degeneration that occurs
in the muscular dystrophies, can be measured noninvasively and
quantitatively in living mice over time. This mouse expresses an
estrogen-responsive Cre-recombinase under the control of the
Pax7 locus and a luciferase reporter gene that is Cre dependent.
Following tamoxifen treatment, luciferase is expressed only in
muscle satellite cells, since these are the only cells in the adult
(other than cells in small regions in the brain) that express Pax7
(7, 8). Therefore, each time the muscle undergoes degeneration
and regeneration, luciferase-expressing satellite cells give rise to
progeny that also express the reporter gene as they proliferate and
differentiate to repair the muscle, and that luciferase activity can
be measured noninvasively in a highly quantitative manner (9–12).
We applied this model to the study of a mouse model of a form of
limb girdle muscular dystrophy (LGMD) and found a remarkable
correlation between the results of noninvasive imaging and dis-
ease activity and progression as determined histopathologically
over the course of 18 months. This technology, which is applica-
ble to all murine models of muscular dystrophy, will dramatically
improve characterizations of the natural history and progression
of muscle diseases and will be an invaluable tool for measuring the
effectiveness of experimental therapeutics.
Characterization of the “regeneration reporter” strain. With the goal
of developing a mouse model to monitor muscle regeneration
as a surrogate for ongoing disease activity in mice with muscu-
lar dystrophies, we used Pax7CreER/LuSEAP mice in which an
estrogen-responsive Cre-recombinase is induced to permanently
activate a luciferase gene in muscle satellite cells (10). To charac-
terize this “regeneration reporter” strain, mice were first imaged
prior to administration of tamoxifen in order to determine the
Conflict of interest: The authors have declared that no conflict of interest exists.
Citation for this article: J Clin Invest. 2013;123(5):2298–2305. doi:10.1172/JCI68458.
Related Commentary, page 1931
The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
association with the increase in cytoplasmic volume and corre-
sponded to a plateau of the luciferase signal determined by biolu-
minescence imaging (Supplemental Figure 3).
These findings in wild-type mice clearly demonstrated the value
of this reporter strain to reflect the proliferative activity of satel-
lite cells and their progeny, providing a noninvasive readout of the
regenerative response in vivo. Furthermore, the specificity of the
luciferase signal for the satellite cell lineage was highly encourag-
ing for the application of this strain to reflect disease activity in
dystrophic muscle. As such, we crossed this reporter strain with a
mouse strain, the SJL strain that is an animal model for the human
disease LGMD2B. LGMD2B is caused by mutations in the dysfer-
lin gene and preferentially affects the proximal limb muscles (13).
There are several mouse models of LGMD2B (14, 15), including
the naturally occurring SJL model (16, 17). As typical of a pro-
gressive muscular dystrophy, histopathological analysis in the SJL
model and in other genetic dysferlinopathy models reveals ongo-
ing degeneration and regeneration of muscle fibers and cumula-
tive fibrosis and adiposis (15, 16, 18, 19).
Monitoring the natural history and progression of disease activity over time.
A hallmark of dystrophic muscle is the degeneration of mature mus-
cle fibers followed by the regeneration of new fibers mediated by
muscle satellite cell progeny (20). We expected that the expression
of luciferase in dystrophic muscle would increase over time, reflect-
ing the regenerative response of the muscle to the ongoing degen-
baseline level of the luciferase signal (Supplemental Figure 1; sup-
plemental material available online with this article; doi:10.1172/
JCI68458DS1). Tamoxifen was then administered for 5 days when
the mice were 2 months of age. Mice were sacrificed 7 days after
the final dose of tamoxifen, and muscles were assessed for the pres-
ence of luciferase-positive satellite cells. Indeed, luciferase-positive
satellite cells were observed in every muscle examined (Figure 1A),
and at the dose of tamoxifen used, an average of 65% of the satel-
lite cells expressed luciferase (Supplemental Figure 2). No other
cells were observed to be luciferase positive, demonstrating the
specificity of the Pax7CreER driver for the satellite cell lineage.
To demonstrate that the proliferative amplification of these cells
could be detected as an increase in luciferase signal, we injured one
of the tibialis anterior (TA) muscles, leaving the contralateral TA
muscle uninjured, in a cohort of mice. Three days after the injury,
the luciferase signals from the injured muscles were dramatically
increased, whereas the signals in the uninjured muscles remained
at baseline levels (Figure 1B). There is always a very low level of
luciferase activity even in uninjured muscles of Pax7CreER/LuSEAP
mice following tamoxifen injection, and this signal is presumably
related to the quiescent satellite cell pool that then expresses luci-
ferase. Histological examination of the injured muscle showed
luciferase-positive myotubes and nascent myofibers during the
process of muscle regeneration (Figure 1C). As myofibers contin-
ued to mature, the intensity of the luciferase signal declined in
The expression of luciferase in resting and injured muscle in Pax7CreER/LuSEAP mice. (A) A luciferase-expressing cell (arrows) residing under
the basal lamina (laminin staining) and expressing the satellite cell marker syndecan-4 1 week after tamoxifen administration to a Pax7CreER/
LuSEAP mouse. Scale bar: 10 μm. (B) Three days after an acute injury to the right TA muscle of a Pax7CreER/LuSEAP mouse, luciferase signals
were detectable only in the injured limb. Scale to the right of the image represents photon emission from the tissue surface and is expressed as
p/s/cm2/sr (or radiance). (C) Luciferase-expressing cells contributed to regenerative myotubes and nascent myofibers during the regenerative
process (days 3–10), but were absent in the uninjured muscle. Scale bar: 50 μm.
2300 The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
cal locations (Figure 3A). Measurements of the luciferase activity
indicated that the proximal limb muscles had significantly more
regeneration than the distal muscles beginning at 6 months of
age (Figure 3B). The luciferase activity in both proximal and distal
muscle groups increased continuously over time, with differences
between them persisting up to 12 months of age (Figure 3B).
Measurement of luciferase expression by histological and biochemical
analyses over time. As a correlate of the changes in luciferase activity
detected by noninvasive imaging and as further validation of the
model, we sacrificed mice at different ages and isolated muscles for
both biochemical and histological studies of luciferase expression.
Based on the temporal progression of luciferase activity determined
by noninvasive imaging (Figure 2C), we expected to see an increase
in the number of luciferase-positive myofibers over time as a result
of the regenerative activity in the muscles. Indeed, we found a sig-
nificant increase in the number of luciferase-positive fibers over
time (Figure 3, C and D), with the proximal limb muscles showing a
greater accretion of luciferase-positive fibers with age than the distal
eration (Figure 2A). Cohorts of 2-month-old wild-type (Dysf+/+) and
dysferlin-deficient (Dysf–/–) Pax7CreER/LuSEAP mice were injected
with tamoxifen to induce luciferase expression in satellite cells. As
additional controls, we injected Dysf–/–/Pax7CreER/LuSEAP mice with
vehicle alone (corn oil) and found no evidence of luciferase expres-
sion (Supplemental Figure 4A). In tamoxifen-treated mice, the luci-
ferase signal increased dramatically over time in hind limb muscles
of the dysferlin-deficient strain, but not in the wild-type strain (Fig-
ure 2, B and C, and Supplemental Figure 4B). This enhanced regen-
erative activity in dysferlin-deficient mice was apparent as early as 3
months of age and increased continuously through 18 months of
age (Figure 2C and Supplemental Figure 4C). Comparisons between
male and female mice showed no significant sex differences with
regard to disease progression (Figure 2D).
Luciferase activity in proximal versus distal limb muscles. LGMD2B
manifests initially and primarily in the proximal limb muscles,
with less involvement of distal limb muscles (21). We assessed
disease activity by bioluminescent signals according to anatomi-
Monitoring disease activity in dysferlin-deficient mice by noninvasive bioluminescence imaging. (A) Diagrammatic representation of the temporal
pattern of luciferase expression in dystrophic Pax7CreER/LuSEAP mice. After tamoxifen administration, a subset of satellite cells express luciferase.
In response to injury or muscle degeneration, these luciferase-positive satellite cells activate and proliferate, giving rise to luciferase-positive myo-
blasts that ultimately differentiate and fuse to form new myofibers, which will then be luciferase positive. (B) Wild-type (left) and dysferlin-deficient
(right) Pax7CreER/LuSEAP mice, injected with tamoxifen at 2 months of age, imaged together at 7 months of age. Scale to the right of the image
represents photon emission from the tissue surface and is expressed as p/s/cm2/sr (or radiance). (C) The luciferase signals from distal hind limb
muscles of both wild-type and dysferlin-deficient mice were measured before tamoxifen administration (Pre-tam), at 3 months of age, and then
monthly up to 18 months of age. P < 0.05 for each measurement between 3 and 18 months of age in the dysferlin-deficient compared with the
control strain; n = 12 Dysf+/+ compared with Dysf–/– at each time point. (D) Luciferase signals from the distal hind limb muscles of male and female
dysferlin-deficient mice (n = 12; same mice as in C).
The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
Noninvasive luciferase imaging in the Dysf–/–/Pax7CreER/LuSEAP
mouse correlates with conventional histopathological manifestations of
disease activity. Classic markers of muscle regeneration in mice
include the appearance of centrally localized nuclei within the
skeletal muscle fibers as well as the appearance of fibers that
express developmental proteins such as embryonic myosin heavy
chain (eMyHc) (22). To validate further the noninvasive imaging
of luciferase activity in Dysf–/–/Pax7CreER/LuSEAP mice as a reflec-
muscles (Figure 4A). Surprisingly, there were relatively few lucifer-
ase-positive fibers in the gastrocnemii (Figure 4A). Measurements of
luciferase activity in whole-muscle extracts also showed an increase
with age (Figure 4B), consistent with the results of noninvasive
imaging. Luciferase-positive fibers were not detected in nondystro-
phic animals, and only background levels of luciferase activity could
be detected in muscles of 12-month-old dysferlin-deficient mice
that were injected with vehicle (Supplemental Figure 5).
Regenerative activity in hind limb muscles of Dysf–/–/Pax7CreER/LuSEAP mice as a function of age. (A) Hind limb muscles of Dysf–/–/Pax7CreER/
LuSEAP mice imaged at 3, 6, 9, and 12 months of age. The luciferase signals increase in the hind limb muscles over time, with more signals
emanating from the proximal muscles after 6 months of age. Scale to the right of the image represents photon emission from the tissue surface
and is expressed as p/s/cm2/sr (or radiance). (B) Luciferase signals were quantified in proximal or distal hind limb muscles of Dysf–/–/Pax7CreER/
LuSEAP mice. **P < 0.05; n = 12. (C) Representative images demonstrating the accrual of luciferase-positive myofibers in quadriceps muscles
of Dysf–/–/Pax7CreER/LuSEAP mice at different ages. Scale bar: 50 μm. (D) Quantitation of the percentage of luciferase-positive quadriceps fibers
from mice sacrificed at 3, 6, 9, and 12 months of age. *P < 0.05; n ≥ 3; value at each time point compared with that at 3 months of age.
2302 The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
sensitively and reliably, we began to monitor the Dysf–/–/Pax7CreER/
LuSEAP mice to assess the changes in luciferase signals over time. We
found that it is possible to monitor disease activity noninvasively,
corroborating our results with histopathological analysis and discov-
ering aspects of the natural history of the SJL mouse model in the pro-
cess. The reliability and reproducibility of this model was extremely
high and comparable to the gold standard of histological measures
of disease activity. Furthermore, individual mice showed patterns of
increases in luciferase activity over time that paralleled the changes
seen in the whole cohort (Supplemental Figure 4B), providing further
support for this imaging system to be used for monitoring disease
activity in different kinds of therapeutic trials in which individual
mice can be used as their own controls in crossover type studies. How-
ever, this technology is not optimal for imaging of muscles, such as
the diaphragm, within the body cavity, as the light signal is attenuated
through layers of tissues and cannot be distinguished from any signal
arising from surface musculature. Finally, it should be noted that the
LuSEAP strain was designed as a dual reporter strain, expressing not
only luciferase but also secreted alkaline phosphatase following Cre-
mediated recombination. However, and as previously noted (10), we
did not find assays of serum alkaline phosphatase levels to provide a
reliable measure of cell number. We thus focused exclusively on the
luciferase reporter for the purpose of these studies.
We specifically chose to use the SJL mouse model for several rea-
sons, several of which are discussed in more detail below. First, it is
a model that exhibits progressive and steady changes over the life-
time of the mouse as opposed to other models of human dystro-
phy, such as the mdx mouse, which shows an early burst of degen-
eration and regeneration followed by a much more stable course
after several months (26–28). Second, there is a longer “preclinical”
and even “prepathological” phase during which we would be able
to assess the sensitivity of our noninvasive imaging methodology
for subtle indications of disease onset as noted above, a finding
that will be critically important in the assessment of therapeutic
interventions. Third, there is an interesting anatomical localiza-
tion of disease severity in the mouse, mimicking that in humans,
which could also be assessed by our noninvasive imaging modal-
ity. Finally, several reports suggest that different pharmacological
interventions may be of benefit in the progressive changes seen in
tion of disease activity, we examined these conventional markers
(Figure 5, A–D). Centrally nucleated myofibers began to appear
at 3 months of age and continuously accrued as the animals aged
(Figure 5, A and B). By 12 months of age, nearly 50% of the fibers
were centrally nucleated in the dysferlin-deficient strain (Figure
5B). Newly regenerated myofibers that express eMyHc could also
be found in increasing numbers as the mice aged (Figure 5, C and
D). In wild-type mice, fewer than 1% of fibers were centrally nucle-
ated, and no eMyHc-positive fibers were detected. Therefore, both
in terms of time course and magnitude of change, noninvasive
imaging faithfully mirrored disease activity and progression by
Here, we present a novel mouse model in which muscle regenera-
tion, a feature of disease activity in muscular dystrophies, can be
followed noninvasively using bioluminescence imaging. By activa-
tion of a satellite cell–specific Cre-recombinase, we marked a major-
ity of muscle satellite cells with the genetic lineage tracer luciferase,
which enabled us to follow muscle regeneration over the course of
18 months in dysferlin-deficient mice. The Pax7CreER strain has been
used previously for lineage tracing of muscle satellite cells in wild-
type mice (9, 10, 12, 23). In one study, postnatal muscle growth, as
assessed by measurement of the incorporation of Pax7-expressing
progenitors into growing myofibers, was monitored for 12 weeks
after tamoxifen administration to wild-type mice at 4 weeks of age
(10). We performed our studies after this postnatal growth period,
at a time when the hind limb muscles have fully matured (24, 25), to
monitor disease activity as a function of regeneration in dystrophic
muscle without any confounding aspects of muscle growth.
We validated the Pax7CreER/LuSEAP mice as a “regeneration
reporter” strain by demonstrating that an acute muscle injury resulted
in a luciferase signal localized only to the injured region of the hind
limb. We observed luciferase-positive myotubes and nascent myofi-
bers with central nuclei, indicating that these newly regenerated fibers
were formed (at least in part) from the progeny of luciferase-express-
ing satellite cells (Figure 1C). In the absence of any injury, luciferase-
expressing myofibers were not found (Figure 1C). After establishing
that this mouse model was capable of reporting regeneration activity
Regenerative activity in proxi-
mal and distal hind limb mus-
cles of Dysf–/–/Pax7CreER/
LuSEAP mice. (A) Represen-
tative images of luciferase-pos-
itive fibers detected in proximal
and distal hind limb muscles of
LuSEAP mice. Scale bar:
50 μm. (B) Quadriceps muscles
of dysferlin-deficient mice have
increasing luciferase enzyme
activity over time. *P < 0.05;
n = 3. Luciferase activities were
measured biochemically in mus-
cles of mice from specific age
groups, as indicated.
The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
regenerative response increased continuously over time, which is
reflective of the progressive nature of LGMD2B (Figure 2C).
Dysferlin deficiency causes a wide range of clinical phenotypes
in humans (13). LGMD2B is generally characterized by the onset
of disease symptoms in the proximal lower limb muscles irrespec-
tive of sex, whereas Miyoshi myopathy (MM), a disease also caused
by mutations in the dysferlin gene, has been described as initially
affecting posterior, distal limb muscles (17, 21). Mouse models of
LGMD2B show that proximal limb muscles are primarily affected
with little disease activity detected in distal muscles (16). In agree-
ment with these findings, we show that the proximal limb muscles
have more regenerative activity than distal muscles beginning at 6
months of age (Figure 3B). Although luciferase signals were greater
in the proximal muscle groups, disease activity was also detected in
the distal limb muscles at 3 months of age (Figure 3B). We found no
significant sex differences in the onset (or progression) of disease
activity in the mice (Figure 2D). Additionally, a greater number of
luciferase-positive myofibers were detected in the TA muscles than
in the gastrocnemii (Figure 4A), a feature that distinguishes between
the 2 diseases, LGMD2B and MM, resulting from mutations in the
dysferlin gene, supporting the conclusion that these mice recapit-
ulate the anatomical distribution of disease activity of LGMD2B.
the SJL mouse (29–31), making it particularly amenable for non-
invasive imaging to be combined with experimental therapeutics.
Clinical manifestations of LGMD2B usually appear during the
second or third decade of life. The rate of progression of this disease
is highly variable among patients, with no correlation between the
age of onset of symptoms and the rate at which the muscles deteri-
orate (32, 33). The SJL mouse has previously been described as hav-
ing a slowly progressive muscular dystrophy by 6 months of age fol-
lowed by rapid disease progression in subsequent months (16, 17).
Likewise, the A/J mouse, another naturally occurring mouse model
of LGMD2B, had virtually no histological evidence of disease activ-
ity until after 5 months of age (14). Our imaging studies revealed
that enhanced regenerative activity began much earlier in the SJL
mouse than had been concluded by the use of conventional markers
of disease activity, demonstrating the enhanced sensitivity of this
imaging technology. By noninvasive imaging, and corroborated by
the detection of luciferase-positive fibers in situ as well as biochem-
ical analyses, we found evidence of disease onset by 3 months of age
(Figure 2C, Figure 3, C and D, and Figure 4B). Although the cumu-
lative luciferase signal was low compared with later time points, the
differences between the SJL and control strains were highly signifi-
cant even at these early time points (Supplemental Figure 4C). This
Histological analyses of muscles
mice over time. (A) Cryosections
of quadriceps muscles of 3-, 6-,
9-, and 12-month-old mice were
stained with H&E to assess for
histopathologic evidence of dis-
ease progression. (B) Quantita-
tive analysis of the percentages
of centrally nucleated myofibers in
quadriceps of Dysf–/–/Pax7CreER/
LuSEAP mice as a function of age.
*P < 0.05, **P < 0.005; n = 3; value
at each time point compared with
that at 3 months of age. (C) Analy-
sis of serial sections from muscles
of mice as in panel A were stained
with an antibody against eMyHc to
assay for regenerating myofibers.
(D) Quantitative analysis of the
percentages of eMyHc-positive
fibers in the quadriceps muscles of
as a function of age. **P < 0.05,
***P < 0.001; n = 3; value at each
time point compared with that at 3
months of age. Scale bars: 50 μm.
2304 The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
were placed on coated glass slides, which were then processed for histolog-
ical or immunohistological staining. Antibodies used in this study were as
follows: chicken anti–syndecan-4 (1:1,000; a gift from Bradley Olwin (Uni-
versity of Colorado, Boulder, Colorado, USA), rabbit anti-laminin (1:1,000;
Sigma-Aldrich L9393), mouse anti-luciferase (1:1,000; Sigma-Aldrich
L2164), and mouse anti-eMyHc (1:100; DSHB clone F1.652-s).
Luciferase biochemical assay. Mice were euthanized as described above, and
muscles were weighed immediately following isolation. The muscles were then
placed into 5 ml of a 2-mg/ml solution of collagenase II for 1 hour at 37°C
with agitation. The muscles were then triturated, and the samples were cen-
trifuged. Supernatants were aspirated, and muscle tissue was lysed as per the
manufacturer’s instructions for the Luciferase Assay System (Promega Inc.).
Isolation of satellite cells. Mice were euthanized and hind limb muscles were
collected. Muscles were resuspended in 0.2% collagenase II and digested for
1 hour and 45 minutes, after which the muscle suspension was triturated
until fully dissociated. The muscles were then subjected to a second diges-
tion in 0.5% collagenase II and 1% dispase for an additional half hour. Sat-
ellite cells were dissociated from the fibers by passing the digested muscle
through a 20-gauge needle. Cells were then collected by centrifugation at
183.7 g and resuspended in 500 μl of Ham’s F-10 medium (Mediatech Inc.).
The cell suspension was incubated with the following primary antibodies:
biotin-labeled VCAM (CD106), FITC-CD45, FITC-CD31, and FITC-Sca1
(1/100; BD Biosciences — Pharmingen) for 45 minutes at 4°C on a shaking
platform. Cells were then washed thoroughly and incubated with anti-FITC
microbeads (1/10; MACS Miltenyi Biotec) for 15 minutes. Cells were then
placed in an AutoMACS cell separation machine according to the manufac-
turer’s instructions (MACS Miltenyi Biotec). Cells that were not FITC posi-
tive were then incubated with anti-biotin microbeads for 15 minutes (1/10;
MACS Miltenyi Biotec) and again subjected to MACS separation according
to the manufacturer’s instructions. The cells were counted and plated on
ECM-coated (1/1,000; Sigma-Aldrich) microscope chamber slides (Lab Tek
II; Fisher Scientific) and incubated at 37°C until fixed.
Statistics. All quantitative data are represented as means, and error bars
indicate the SEM. Data were subjected to either 2-tailed Mann-Whitney
test (for disease progression analysis comparing wild-type to dysferlin-de-
ficient mice) or 2-tailed Student’s t tests (when comparing male and female
imaging data, proximal and distal hind limb, and quantitations from his-
tological analyses) to determine the significance of calculated differences.
These differences were considered statistically significant at P < 0.05.
Study approval. Animals were handled and housed according to the guide-
lines set forth by the Veterinary Medical Unit of the VA Palo Alto Health
Care System, and all procedures were preapproved by the IACUC prior to
We would like to acknowledge Charles Keller for his seminal con-
tributions to the field in terms of the generation of the transgenic
mouse strains that were used in these studies and that we and
others have found to be so valuable in studies of muscle stem cell
biology and muscle disease studies. This work was supported by
grants from the Jain Foundation and the NIH (Director’s Pioneer
Award DP1 OD000392) to T.A. Rando.
Received for publication December 19, 2012, and accepted in
revised form February 21, 2013.
Address correspondence to: Thomas A. Rando, Department of
Neurology and Neurological Sciences, Stanford University School
of Medicine, Stanford, California 94305-5235, USA. Phone:
650.849.0199; Fax: 650.858.3935; E-mail: firstname.lastname@example.org.
Previously, it was reported that disease activity in the SJL mouse
model could be imaged by MRI using the enhanced contrast-
ing agent Gadofluorine-M (34). While it was possible to detect
increased permeability of muscle fibers to this agent, and this
correlated with advanced disease progression, it was not sensi-
tive enough to detect early pathologic changes. Additionally, the
detection limit was dependent upon the amount of contrasting
agent administered to the mice, which may have been limiting in
the analysis (34). Although MRI does not require breeding of dys-
trophic mice with reporter mice, noninvasive imaging of disease
activity using luciferase bioluminescence is less expensive, more
widely available, and much more versatile for laboratory studies
to monitor disease progression noninvasively for natural history
studies or studies of response to experimental therapeutics.
The goal of this study was to generate a mouse model in which
muscle regenerative activity could be measured over time to mon-
itor the progression of disease activity or response to therapy for
muscular dystrophies. Traditionally, the efficacy of drug treat-
ments for the muscular dystrophies has been assayed using, alone
or in combination, strength measurements, serum-biomarker
concentrations, and changes in histopathology. As an alternative,
the Pax7CreER/LuSEAP system will be invaluable for testing phar-
macological, cellular, and genetic treatments for muscular dys-
trophies. Increased sensitivity, reliable quantitation, the ability to
use each animal as its own control, and obviating the need to do
labor- and time-intensive histopathology all will allow for more
high-throughput analysis of treatments in vivo with this noninva-
sive method to monitor disease progression.
Animals. The Pax7CreER and LuSEAP mice were a gift of Charles Keller, Oregon
Health and Science University, Portland, Oregon, USA. SJL mice were pur-
chased from Jackson Laboratories.
Bioluminescence imaging. Bioluminescent imaging was performed using
the Xenogen IVIS-Spectrum System (Caliper Life Sciences). Mice were anes-
thetized using 2% isoflurane and 100% oxygen at a flow rate of 2.5 l/min.
Then 300 μl of a 50 mg/ml sterile D-Luciferin firefly substrate (Biosynth
International Inc.) dissolved in PBS was administered by i.p. injection, and
23 minutes after substrate injection, the mice were imaged for 30 seconds
at the maximal light collection (f-stop 1) at the highest resolution (small
binning). Each image was saved for subsequent analysis. The scales to the
right of images in Figure 1B, Figure 2B, and Figure 3A represent the pho-
ton emission from the tissue surfaces and are expressed as photons/sec-
ond/centimeter squared/steradian (p/s/cm2/sr) (or radiance).
Image analysis. Analysis of each image was performed using Living Image
Software, version 4.0 (Caliper Life Sciences). Briefly, a manually-generated cir-
cle was placed on top of the region of interest (ROI) and resized to completely
surround the limb or the specified region on the wild-type mouse. This ROI
outline was then duplicated and placed over the limb of the dysferlin-defi-
cient mouse. In cases where a control wild-type mouse was not imaged simul-
taneously, the ROI was encircled only for the experimental mouse.
Skeletal muscle injury. Mice were anesthetized using 2% isoflurane and 100%
oxygen at a flow rate of 2.5 l/min. The hind limbs were shaved, and 25 μl
of 1.2% BaCl2 was injected into TA muscles using 10-cc insulin syringes (9).
Histology and immunohistochemistry. At the specified time points, mice were
euthanized using CO2 asphyxiation followed by cervical dislocation. Hind
limb muscles were submerged in 0.5% PFA for 5 hours and then dehydrated
in 20% sucrose overnight at 4°C. The muscles were then submerged in
OCT (Sakura Finetek USA Inc.) and frozen for approximately 30 seconds
in liquid nitrogen chilled in isopentane. Muscle cryosections (8-μm-thick)
The Journal of Clinical Investigation http://www.jci.org Volume 123 Number 5 May 2013
1. Cohn RD, Campbell KP. Molecular basis of muscular
dystrophies. Muscle Nerve. 2000;23(10):1456–1471.
2. Goyenvalle A, Seto JT, Davies KE, Chamberlain J.
Therapeutic approaches to muscular dystrophy.
Hum Mol Genet. 2011;20(R1):R69–R78.
3. Vainzof M, et al. Animal models for genetic neuro-
muscular diseases. J Mol Neurosci. 2008;34(3):241–248.
4. Lieberman JS, Taylor RG, Fowler WM. Serum crea-
tine phosphokinase variations in dystrophic mice.
Exp Neurol. 1981;73(3):716–724.
5. Dellorusso C, Crawford RW, Chamberlain JS,
Brooks SV. Tibialis anterior muscles in mdx mice
are highly susceptible to contraction-induced
injury. J Muscle Res Cell Motil. 2001;22(5):467–475.
6. McIntosh LM, Baker RE, Anderson JE. Magnetic reso-
nance imaging of regenerating and dystrophic mouse
muscle. Biochem Cell Biol. 1998;76(2–3):532–541.
7. Seale P, Sabourin LA, Girgis-Gabardo A, Mansouri
A, Gruss P, Rudnicki MA. Pax7 is required for
the specification of myogenic satellite cells. Cell.
8. Buckingham M, et al. Myogenic progenitor cells
in the mouse embryo are marked by the expres-
sion of Pax3/7 genes that regulate their survival
and myogenic potential. Anat Embryol (Berl).
9. Brack AS, et al. Increased Wnt signaling during
aging alters muscle stem cell fate and increases
fibrosis. Science. 2007;317(5839):807–810.
10. Nishijo K, et al. Biomarker system for studying
muscle, stem cells, and cancer in vivo. FASEB J.
11. Quach NL, Biressi S, Reichardt LF, Keller C, Rando
TA. Focal adhesion kinase signaling regulates the
expression of caveolin 3 and beta1 integrin, genes
essential for normal myoblast fusion. Mol Biol Cell.
12. Bjornson CR, Cheung TH, Liu L, Tripathi PV,
Steeper KM, Rando TA. Notch signaling is neces-
sary to maintain quiescence in adult muscle stem
cells. Stem Cells. 2012;30(2):232–242.
13. Amato AA, Brown RH. Dysferlinopathies. Handb
Clin Neurol. 2011;101:111–118.
14. Ho M, et al. Disruption of muscle membrane and
phenotype divergence in two novel mouse mod-
els of dysferlin deficiency. Hum Mol Genet. 2004;
15. Bansal D, et al. Defective membrane repair in
dysferlin-deficient muscular dystrophy. Nature.
16. Bittner RE, et al. Dysferlin deletion in SJL mice (SJL-
Dysf) defines a natural model for limb girdle muscu-
lar dystrophy 2B. Nat Genet. 1999;23(2):141–142.
17. Weller AH, Magliato SA, Bell KP, Rosenberg NL.
Spontaneous myopathy in the SJL/J mouse:
pathology and strength loss. Muscle Nerve.
18. Nemoto H, Konno S, Nakazora H, Miura H, Kuri-
hara T. Histological and immunohistological
changes of the skeletal muscles in older SJL/J mice.
Eur Neurol. 2007;57(1):19–25.
19. Kobayashi K, Izawa T, Kuwamura M, Yamate J. The
distribution and characterization of skeletal mus-
cle lesions in dysferlin-deficient SJL and A/J mice.
Exp Toxicol Pathol. 2010;62(5):509–517.
20. Wallace GQ, McNally EM. Mechanisms of muscle
degeneration, regeneration, and repair in the mus-
cular dystrophies. Annu Rev Physiol. 2009;71:37–57.
21. Bushby KM. Dysferlin and muscular dystrophy.
Acta Neurol Belg. 2000;100(3):142–145.
22. Reznik M, Engel WK. Ultrastructural and his-
tochemical correlations of experimental muscle
regeneration. J Neurol Sci. 1970;11(2):167–185.
23. Lepper C, Fan CM. Inducible lineage tracing of
Pax7-descendant cells reveals embryonic origin of
adult satellite cells. Genesis. 2010;48(7):424–436.
24. Schultz E. Satellite cell proliferative compart-
ments in growing skeletal muscles. Dev Biol.
25. White RB, Bierinx AS, Gnocchi VF, Zammit PS.
Dynamics of muscle fibre growth during postnatal
mouse development. BMC Dev Biol. 2010;10:21.
26. Bulfield G, Siller WG, Wight PA, Moore KJ. X
chromosome-linked muscular dystrophy (mdx)
in the mouse. Proc Natl Acad Sci U S A. 1984;
27. Dangain J, Vrbova G. Muscle development in mdx
mutant mice. Muscle Nerve. 1984;7(9):700–704.
28. DiMario JX, Uzman A, Strohman RC. Fiber regener-
ation is not persistent in dystrophic (MDX) mouse
skeletal muscle. Dev Biol. 1991;148(1):314–321.
29. Suzuki N, et al. Continuous administration of
poloxamer 188 reduces overload-induced muscu-
lar atrophy in dysferlin-deficient SJL mice. Neurosci
30. Nemoto H, et al. Anti-TNF therapy using etaner-
cept suppresses degenerative and inflammatory
changes in skeletal muscle of older SJL/J mice. Exp
Mol Pathol. 2011;90(3):264–270.
31. Rayavarapu S, Van der Meulen JH, Gordish-Dress-
man H, Hoffman EP, Nagaraju K, Knoblach SM.
Characterization of dysferlin deficient SJL/J
mice to assess preclinical drug efficacy: fasudil
exacerbates muscle disease phenotype. PLoS One.
32. Paradas C, et al. Redefining dysferlinopathy pheno-
types based on clinical findings and muscle imag-
ing studies. Neurology. 2010;75(4):316–323.
33. Gayathri N, et al. Dysferlinopathy: spectrum of
pathological changes in skeletal muscle tissue.
Indian J Pathol Microbiol. 2011;54(2):350–354.
34. Schmidt S, et al. Dysferlin-deficient muscular dys-
trophy: gadofluorine M suitability at MR imaging
in a mouse model. Radiology. 2009;250(1):87–94.