Download A mutation in α-tropomyosin affects muscle strength, maturation and

© 2001 Oxford University Press
Human Molecular Genetics, 2001, Vol. 10, No. 4
317–328
ARTICLE
A mutation in α-tropomyosinslow affects muscle
strength, maturation and hypertrophy in a mouse
model for nemaline myopathy
Mark A. Corbett1, C. Stephen Robinson1, Greta F. Dunglison1, Nan Yang2,
Josephine E. Joya1, Angus W. Stewart5, Christina Schnell2, Peter W. Gunning3,4,
Kathryn N. North2,4 and Edna C. Hardeman1,+
1Muscle
Development Unit, Children’s Medical Research Institute, Locked Bag 23, Wentworthville, New South Wales
2145, Australia, 2Neurogenetics Research Unit and 3Oncology Research Unit, The New Children’s Hospital, PO Box
3515, Parramatta, New South Wales 2124, Australia, 4Department of Pediatrics and Child Health, University of
Sydney, Sydney, New South Wales 2006, Australia and 5School of Biomedical and Sports Science, Edith Cowan
University, 100 Joondalup Drive, Joondalup, Western Australia 6027, Australia
Received 4 August 2000; Revised and Accepted 12 December 2000
Nemaline myopathy is a hereditary disease of skeletal muscle defined by a distinct pathology of electrondense accumulations within the sarcomeric units called rods, muscle weakness and, in most cases, a slow
oxidative (type 1) fiber predominance. We generated a transgenic mouse model to study this disorder by
expressing an autosomal dominant mutant of α-tropomyosinslow previously identified in a human cohort. Rods
were found in all muscles, but to varying extents which did not correlate with the amount of mutant protein
present. In addition, a pathological feature not commonly associated with this disorder, cytoplasmic bodies,
was found in the mouse and subsequently identified in human samples. Muscle weakness is a major feature
of this disease and was examined with respect to fiber composition, degree of rod-containing fibers, fiber
mechanics and fiber diameter. Hypertrophy of fast, glycolytic (type 2B) fibers was apparent at 2 months of
age. Muscle weakness was apparent in mice at 5–6 months of age, mimicking the late onset observed in
humans with this mutation. The late onset did not correlate with observed changes in fiber type and rod
pathology. Rather, the onset of muscle weakness correlates with an age-related decrease in fiber diameter
and suggests that early onset is prevented by hypertrophy of fast, glycolytic fibers. We suggest that the clinical
phenotype is precipitated by a failure of the hypertrophy to persist and therefore compensate for muscle
weakness.
INTRODUCTION
Nemaline myopathy is a hereditary disease of skeletal muscle
defined by a distinct pathology of electron-dense accumulations within the sarcomeric units called rods. Nemaline myopathy
has been classified by the ENMC International Consortium on
Nemaline Myopathy into six subcategories based on clinical
manifestations of the disorder, including age at onset and
severity of weakness: severe congenital, typical, intermediate
congenital, mild form of childhood- or juvenile-onset, adultonset and other. Muscle weakness associated with this disorder
can be either static or slowly progressive and, in most cases, a
slow oxidative (type 1) fiber predominance as well as atrophy
of type 1 fibers is observed (reviewed in ref. 1). Studies on
human biopsy samples have proposed either conversion from
+To
type 2 to type 1 fibers (2) or a disruption of normal fiber development (3) as mechanisms for this process.
Nemaline rods are composed primarily of the Z-line protein
α-actinin (4,5). Ultrastructural studies also have suggested that
sarcomeric α-actin is a component (6,7; C.S. Robinson,
personal communication) and the intermediate filament
protein desmin can be found in the periphery of areas where
rod disruption occurs (4,5). Rods appear to form as extensions
of the Z-line with which they share structural similarities (8).
They may also occur in association with central cores (core–rod
myopathy) (9), as a secondary phenomenon in mitochondrial
disorders (9), in patients with HIV (10,11) as well as in experimental conditions such as tenotomy in the rat (12). Nemaline
rods have also been observed in the dog (13) and cat (14) as
naturally occurring disease models. The significance of rods in
whom correspondence should be addressed. Tel: +61 2 9687 2800; Fax: +61 2 9687 2120; Email: [email protected]
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Human Molecular Genetics, 2001, Vol. 10, No. 4
relation to muscle weakness is unknown. Sequential analyses
of muscle biopsies from patients with nemaline myopathy
suggest that there is no correlation between the number of
fibers containing rods and the degree of muscle weakness (15).
The composition of rods and their location at the site of
Z-line and thin filament association suggested that candidate
genes for nemaline myopathy are likely to be involved with the
thin filament network. Disease-associated mutations have been
identified in humans in three genes encoding sarcomeric thin
filament proteins: αTmslow (16,17), nebulin (18) and α-skeletal
actin (19). α-actinin is also a candidate gene as the formation
of rods has been demonstrated in hemizygous Drosophila
knockouts (20) and cell culture with mutant forms of the
protein (21). Laing et al. (17) identified the first diseaseassociated mutation for nemaline myopathy in a large
Australian pedigree diagnosed with childhood-onset nemaline
myopathy. The mutation was a single base change which
converted a methionine to arginine at codon 9 (Met9Arg) in the
gene encoding αTmslow (TPM3), likely resulting in a dominant
negative mutation. A second mutation with recessive inheritance
has also been identified in αTmslow in association with nemaline
myopathy (16).
The tropomyosins comprise a large family of structural
proteins. In skeletal muscle, tropomyosin forms a thin
filament through the head-to-tail polymerization of rod-like
heterodimers of α and β subunits (22). The tropomyosin filament
lies in the major groove of filamentous α-actin and plays a
regulatory role in muscle contraction which is dependent on
Ca2+ ion concentration (23). The head and tail domains of the
protein are believed to have a function in both dimerization and
actin binding (22). The Met9Arg missense mutation is in a
well-conserved region of the N-terminal portion of the molecule
that may be essential for actin binding. It has been postulated
that the addition of another basic residue to this already basic
rich region may strengthen actin binding (17). Alternatively, a
reduction in the affinity between tropomyosin and actin could
allow fortuitous interactions between α-actinin and actin
resulting in the extension of α-actinin into the thin filament. In
striated muscle there exists a tight regulation of gene expression
between related family members (24). A mutation in any one
of the skeletal muscle tropomyosins therefore has the potential
to cause serious disruption to the sarcomere.
We expressed the dominant negative αTmslow(Met9Arg)
mutation specifically in skeletal muscle of mice in order to
generate a mouse model for this disease. All features of the
disease observed in the human cohort are present in the mouse
which allowed us to address key issues concerning the mechanism
of type I fiber predominance, rod accumulation in fibers and
onset of muscle weakness. The increase in slow, oxidative
fibers was established at 1 month of age and was not progressive,
indicating impaired fiber maturation rather than fiber conversion
as a basis for this aspect of the disease. Differences were
observed in a number of different parameters in the absence of
overt muscle weakness. Nemaline fibers displayed altered
strontium sensitivity suggesting an alteration in the tropomyosin/
troponin regulatory complex. The percentage of rodcontaining fibers varied significantly between different
muscles, did not correlate with amount of mutant protein
present and increased gradually between 2 and 12 months.
Hypertrophy of fast, glycolytic (2B) fibers was apparent at
2 months of age and it is the age-related decline in this hyper-
Figure 1. Transgene constructs. (A) TnIslow–αTmslow(Met9Arg) construct uses
the human troponin Islow promoter (striped) to drive expression of the
αTmslow(Met9Arg) mutant cDNA (black) with an SV40 3′-UTR sequence
(hatched) specifically in slow fiber types. (B) HSA –αTmslow(Met9Arg) and
(C) HSA–αTmslow(wt) use the human skeletal actin promoter (gray) to drive
expression of the αTmslow(Met9Arg) mutant or the wild-type (white) cDNA
sequence predominantly in fast fiber types.
trophy that correlates with the onset of muscle weakness which
occurs at 5–6 months of age.
RESULTS
Generation of mice expressing the αTmslow(Met9Arg)
transgene
We have taken two approaches to generate a mouse model for
early childhood-onset nemaline myopathy. Both take
advantage of the autoregulatory properties of the sarcomeric
tropomyosin gene family. This avoids the problem of overexpression of the introduced protein because downregulation
of the endogenous product compensates to maintain a constant
pool size (24). In the first approach, we imitated the slow fiberspecific expression of the mutant protein. In order to achieve
this, we linked the coding region for αTmslow(Met9Arg) to a
region of the human troponin I slow gene that drives
expression specifically in slow-twitch (type 1) muscle fibers
(25). Six founder lines were generated with this construct
designated TnIslow–αTmslow(Met9Arg) (Fig. 1A). The second
approach took into consideration the significant difference in
fiber composition of muscles between mouse and human. The
majority of mouse muscles are composed of ∼90% or greater
fast-twitch (type 2) fibers (26,27). In contrast, the majority of
muscles in humans contain ∼40% or greater type 1 fibers.
Therefore, to achieve a mouse model in which a significant
number of fibers would be affected by the presence of the
mutant protein, we expressed αTmslow(Met9Arg) in type 2
fibers of the mouse. In this construct, the coding region of
αTmslow(Met9Arg) was linked to the human skeletal actin
(HSA) promoter that drives expression preferentially in 2B
fibers of the mouse (28; E.C. Hardeman, unpublished data).
Five founder lines were generated with this construct designated HSA–αTmslow(Met9Arg) (Fig. 1B). To establish that any
pathological features arising in the HSA–αTmslow(Met9Arg)
Human Molecular Genetics, 2001, Vol. 10, No. 4
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mice were due to the expression of the mutant protein and not
the presence of a slow Tm protein in a fast-twitch fiber, a third
construct was made. The coding region of the wild-type
αTmslow protein was linked to the HSA promoter region. Nine
founder lines were generated with this construct designated
HSA–αTmslow(wt) (Fig. 1C). Transgenic mice of all lines
were fertile and a 50% transmission frequency of the transgene
was observed in all lines. All of the analyses described in the
subsequent sections were performed on the TnIslow–
αTmslow(Met9Arg), HSA–αTmslow(Met9Arg) and HSA–
αTmslow(wt) lines of mice unless otherwise indicated.
Mice expressing αTmslow(Met9Arg) develop nemaline rods
Nemaline rods were present in the slow fiber-containing soleus
and complex zone of the crural muscle block in one transgenic
line expressing the TnIslow–αTmslow(Met9Arg) construct and in
all fast fiber-containing muscles of three lines expressing the
HSA–αTmslow(Met9Arg) construct (discussed below). Nemaline
rods were observed in both the center and the periphery of
muscle fibers. Rod length was between 2 and 3 µm in the
muscles examined. No rods were observed in lines expressing
HSA–αTmslow(wt) which established that inappropriate
expression of αTmslow in fast fibers does not contribute to rod
pathology. Rods were commonly observed in clusters and
areas surrounding the Z-line, resulting in Z-line streaming and
disruption of the sarcomeric register (Fig. 2A). The nemaline
rods are comparable to those seen in patients who have the
Met9Arg mutation in TPM3 (Fig. 2B), and appear to form as
lateral extensions of the Z-line. The identity of the rods
was confirmed by indirect immunofluorescent staining of
longitudinal sections of plantaris for α-actinin 2 (Fig. 3), a
major component of nemaline rods and the Z-line (4,5). This
result unambiguously demonstrates that the αTmslow(Met9Arg)
mutation is responsible for rod pathology in humans with this
TPM3 mutation and that it acts through a dominant mechanism
(17).
Mice expressing αTmslow(Met9Arg) develop cytoplasmic
bodies and tubular aggregates of the sarcoplasmic
reticulum
Two inclusions not typically associated with nemaline
myopathy in humans were observed only in the HSA–
αTmslow(Met9Arg) lines and only in one muscle, the superficial
gastrocnemius (SG). Cytoplasmic bodies were present from
2 months of age at a frequency of 3.5–10% of fibers (Fig. 4A).
The cytoplasmic bodies have a different electron density and
are larger than nemaline rods (∼5 µm in diameter). These
structures were observed in areas where complete breakdown
of the sarcomeric structure had occurred and were sometimes,
but not always, associated with clusters of nemaline rods.
Further analysis of a patient with the Met9Arg TPM3 mutation
has revealed features similar to cytoplasmic bodies in the
deltoid muscle (Fig. 4B). Tubular aggregates of the sarcoplasmic reticulum appeared in the SG of transgenic mice from
1 month of age and the percentage of affected fibers reached
45% by 12 months of age (Fig. 4C and D). Tubular aggregates
have been observed in cases of severe muscle cramps and
periodic paralysis in humans (29,30).
Figure 2. Nemaline rods in HSA–α Tmslow(Met9Arg) transgenic and human
(Met9Arg) muscle. Electron micrograph of nemaline rods in longitudinal
sections from (A) the EDL muscle of 2-month-old HSA–αTm slow(Met9Arg)
line 4 (20 100×; scale bar, 1 µm) and (B) the left deltoid muscle of a human
with childhood-onset αTm slow(Met9Arg) nemaline myopathy (4000×; scale
bar, 5 µm). Arrowheads indicate areas of Z-line streaming.
Rod formation correlates with a threshold level of
αTmslow(Met9Arg) mRNA
Rod pathology was observed in those lines of mice in which a
level of αTmslow(Met9Arg) mRNA approximating that of the
endogenous αTmslow was achieved. αTmslow(Met9Arg)
transcript levels were determined in the crural muscle block
(without soleus) of HSA–αTmslow(Met9Arg) lines. The three
lines that developed rod pathology expressed the mutant Tm
transcript at levels of ∼160% (line 4), 120% (line 14) and 70%
(line 9) of that of the endogenous αTmslow gene in slow fibers
(Fig. 5). Further support for this threshold phenomenon was
observed in the highest expressing TnIslow–αTmslow(Met9Arg)
line 49. When the transgenic locus in this line is in the
hemizygous state, the αTmslow(Met9Arg) transcript level is
∼50% of the endogenous αTmslow in slow fibers and rods are
absent. However, when bred to homozygosity to increase the
transgene expression level, rods develop in the soleus muscles
of these mice. Thus, expression from the αTmslow(Met9Arg)
locus must approach levels comparable to the endogenous
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Figure 3. Confocal detection of α-actinin 2 showing nemaline rods as Z-line
extensions. Longitudinal section (100 µm) of EDL muscle from 6-month-old
HSA–αTmslow(Met9Arg) line 4 labeled with α-actinin 2 antibody (4B2)
visualized with FITC. The open arrow indicates Z-line; the closed arrow
indicates Z-line extension. Scale bar, 10 µm.
αTm loci in order for rods to form. This is analogous to the
mutant to normal αTmslow ratio found in the human condition.
The number of fibers containing rods and rod size are
different in different muscles
The HSA–αTmslow(Met9Arg) lines show a high percentage of
fibers containing rods in the diaphragm (49%) and forearm
muscles [70% in the extensor carpi ulnaris (ECU)] compared
with much lower numbers in the SG (2%) and tibialis anterior
(TA) (17 %) (Fig. 6A). Differential rod formation in different
fiber types cannot account for this phenomenon since these
muscles have the same fiber composition [3–5% type I, 97–95%
type 2 (data not shown)]. This difference in percentage of rodcontaining fibers between muscles is not a result of differential
expression of the mutant protein. The amount of mutant
protein expressed in these muscles is shown on an isoelectric
focusing western blot for comparison (Fig. 6B). The mutant
protein can be clearly seen in all lines as the upper (most basic)
band in the transgenic mouse muscle extracts. α-tropomyosinfast
(αTmfast) and β-tropomyosin (βTm) run as a doublet as the
lower (most acidic) band. In slow muscle (soleus) extracts, the
band running below the mutant and above the αTmfast and βTm
bands is the αTmslow band. The relative amount of mutant
tropomyosin in the various muscles normalized to endogenous
actin is shown in Figure 6C. Our data show that, although a
threshold level of protein is required for rod formation to
occur, the level of protein expression in each muscle does not
ultimately define the degree of rod pathology. A similar analysis
was not possible in TnIslow–αTmslow(Met9Arg) line 49 mice
since only two muscles in this line of mice contain rods.
There is an increase in the number of rod-containing fibers in
the extensor digitorum longus (EDL) between 2 and 12 months
(Fig. 6D). Increases in the number of rod-affected fibers with
age have been observed in some, but not all, human follow-up
studies (15). We also measured the mean size of the rods in
selected muscles, EDL, diaphragm and ECU at 6 months of
Figure 4. Cytoplasmic bodies and tubular aggregates in HSA–αTmslow(Met9Arg) transgenic muscle. Electron micrographs of (A) cytoplasmic body (8000×; scale
bar, 5 µm) and (C and D) tubular aggregates of the sarcoplasmic reticulum [(C) 77 000×; scale bar; 100 nm; (D) 15 000×; scale bar, 1 µm) in longitudinal sections
(70 nm) of SG muscle of 2-month-old HSA–αTm slow(Met9Arg) transgenic line 4. (B) Cytoplasmic body in the left deltoid muscle of a human with childhood-onset
αTmslow(Met9Arg) nemaline myopathy (12 000×; scale bar, 5 µ m).
Human Molecular Genetics, 2001, Vol. 10, No. 4
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Skinned fibers from nemaline mice show minor changes in
response to Sr2+
Figure 5. Northern blot analysis of αTmslow(Met9Arg) transgene and endogenous αTmslow transcripts. Total RNA (10 µ g) was analyzed from (A) the crural
muscles (without soleus) of the HSA–αTmslow(Met9Arg) lines 4, 9, 14, 20 and
33 or (B) the soleus muscle of the TnIslow–α Tmslow(Met9Arg) lines 1, 22, 31,
41 and 49. Transgene (2.4 kb) and endogenous (1.3 kb) transcripts were
detected with a probe to the first 180 bases of coding sequence for αTmslow.
Loading differences are indicated by detection of the 18S rRNA on the same
blot (18S). C, non-transgenic crural (without soleus); S, non-transgenic soleus.
age. Rods were significantly longer in the ECU (P < 0.001)
than in the EDL and the diaphragm (Fig. 6E). The size of rods
was within the size range recorded for cases of the human
disease (1). Rods did not increase in length in the EDL between
2 and 12 months.
Nemaline mice have an increased number of slow/fast
oxidative fibers
Slow or type 1 fiber predominance is frequently observed in
human nemaline myopathy (1). This could result from an
active conversion of type 2 fibers to type 1 fibers, a disruption
in the normal developmental program of fiber maturation, or a
combination of these mechanisms. Oxidative metabolism is
more energy efficient and an increase in slow/fast oxidative
fibers (types 1 and 2A) may be a natural mechanism to combat
muscle weakness. The EDL muscles of HSA–αTmslow(Met9Arg)
lines were examined at 1, 2, 6 and 12 months for fiber type
composition as defined by the presence of myosin heavy chain
isoforms. From 2 months of age, the EDLs of the transgenics
contain a higher percentage of slow/fast oxidative fibers than
their wild-type littermates (Fig. 7). Between 1 and 12 months,
the fiber type composition in the EDL of HSA–αTmslow(Met9Arg)
lines remained relatively static, suggesting a disruption to
normal maturation. A parallel analysis of HSA–αTmslow(wt)
transgenic mice with similar levels of transgene expression
showed normal fiber composition and maturation. This
confirms that the increase in slow/fast oxidative fibers
observed in HSA–αTmslow(Met9Arg) lines is due to the
Met9Arg mutation and not the inappropriate expression of the
slow Tm isoform in fast fibers. Fiber composition in the soleus
of TnIslow–αTmslow(Met9Arg) line 49 mice was similar to wildtype. This may indicate that the normal fiber composition of
the soleus is at the endpoint of the slow/oxidative composition
achievable in a mouse maintained in a laboratory environment.
Single fiber mechanical studies were performed on 15 fibers
from HSA–αTmslow(Met9Arg) line 4 mice and 23 fibers from
HSA–αTmslow(wt) line 41 mice to determine whether aspects
of fiber performance could be attributed to the mutant Tm. The
fibers fell into two distinct populations based on the pCa50–pSr50
differences (31). Values of >1.4 were designated population 1;
whereas, values <1.4 were designated population 2. Based on
this delineation, Table 1 shows the calculated mean of each
parameter. The population 2 fibers yielded significant changes
in the nSr and pSr10 values; however, there were no significant
changes in any parameter for population 1 fibers. No slow
force oscillations were noted in any of the fibers tested. The
only difference observed between nemaline and control population 2 fibers was in strontium sensitivity, which could
suggest a difference in the functionality of the regulatory
protein complex.
Mice expressing Tmslow(Met9Arg) develop late-onset
muscle weakness
Mice from the various transgenic lines did not exhibit an overt
clinical phenotype; however, this was not unexpected since
humans expressing the Tmslow(Met9Arg) mutation display a
mild muscle weakness. Since a test of gross muscle strength,
such as the Gower’s maneuver, is used to diagnose muscle
weakness in humans, we used a standard whole-body strength
and fatigability test to assess gross muscle strength in the mice.
Mice from HSA–αTmslow(Met9Arg) line 4 were analyzed
since this test is most effective if a significant number of
muscles are affected. Mice hanging by their forepaws were
required to pull themselves up onto a metal bar in a series of
15 consecutive tests (32) (Fig. 8). Between 2 and 4 months of
age, no significant differences in the average pass rate over the
15 tests were observed in HSA–αTmslow(Met9Arg) line 4 mice
in comparison with nontransgenic littermates. At 5–6 months
of age, HSA–αTmslow(Met9Arg) transgenic mice exhibited a
reduced capacity to perform this test (control 72 ± 8%, transgenic 55 ± 10%). This trend continued, with a further drop in
the performance of the nemaline mice between 10 and
13 months of age. The average pass rate of the nemaline mice
was 37 ± 7% in comparison with 66 ± 8 % for the nontransgenic littermates (P < 0.05). The late onset of muscle
weakness is reminiscent of the childhood-onset disorder
observed in humans with the αTmslow(Met9Arg) mutation.
Mice expressing αTmslow(Met9Arg) display hypertrophy of
fast, glycolytic fibers
In some human nemaline myopathy patients, a compensatory
hypertrophy of unaffected fibers is associated with less severe
weakness (15). We measured fiber diameters in the EDL of
HSA–αTmslow(Met9Arg) mice at 2, 6 and 12 months of age to
determine whether hypertrophy is a feature in this mouse
model. Slow/fast, oxidative fiber types (types 1 and 2A) have
smaller diameters than the fast, glycolytic fibers (types 2X and
2B). Since the HSA–αTmslow(Met9Arg) lines have a greater
number of oxidative fiber types, we grouped 1/2A and 2X/2B
fibers to avoid a bias in our measurements towards atrophied
fibers. At all ages we observed that the mean diameter of 2X/
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Figure 6. Properties of rods in muscles of HSA–α Tmslow(Met9Arg) lines.
(A) Rods were detected in toluidine blue-stained longitudinal sections (0.3 µm)
of resin-embedded extensor digitorum lonus (EDL), superficial gastrocnemius
(SG), tibialis anterior (TA), diaphragm (DIA), flexor carpi radialis (FCR),
flexor digitorum profundus (FDP), flexor carpi ulnaris (FCU) and extensor
carpi ulnaris (ECU) muscles from 2-month-old HSA–αTmslow(Met9Arg) line 4.
Rod-containing fibers are expressed as the mean percentage of the total number
of fibers examined in one muscle from three to five individuals (± SEM).
(B) Protein extracts (15 µg) of EDL, SG, TA, soleus (SOL), DIA, FCR, FDP,
FCU and ECU muscles from 2-month-old HSA –αTmslow(Met9Arg) line 4
were analyzed by isoelectric focusing western blots. αTmslow(Met9Arg) (band
indicated by Met9Arg) and endogenous sarcomeric Tm proteins, αTmslow,
αTmfast and βTm (bands indicated by α slow, α fast and β, respectively) were
detected with a sarcomeric Tm antibody (CH1). Loading differences were
determined using an antibody (C4) that recognizes both sarcomeric (α -sarc)
and cytoplasmic (β,γ) actins. +, transgenic muscle; –, non-transgenic littermate muscle. (C) Densitometry values of the Met9Arg levels shown in (B)
normalized to actin. (D) Progression of rod pathogenesis in HSA –
αTmslow(Met9Arg) EDL. Rods were detected as described in (A) from EDL of
2-, 6- and 12-month-old HSA–αTmslow(Met9Arg) lines 4 and 14. Rod-containing fibers are expressed as the mean percentage of the total number of fibers
examined in one muscle from four individuals per time point (± SEM). (E) Rod
lengths in different HSA–αTmslow(Met9Arg) muscles. Rods were detected as
described in (A) from EDL, ECU and diaphragm of 6-month-old HSA –
αTmslow(Met9Arg) line 4. Rod dimensions were measured from electron
micrographs. At least 85 rods were measured for each muscle (± SEM). The
size increase for ECU (*) is significant (P < 0.001) as determined by singlefactor ANOVA.
Similar changes in fiber diameter mediated through a natural
program of muscle maturation in human may also be an important determinant for age at onset of muscle weakness.
DISCUSSION
2B fibers is greater in HSA–αTmslow(Met9Arg) lines (Fig. 9).
No alteration was observed in the diameter of 1/2A fibers.
Indeed, there appeared to be an age-related increase in the
mean 2X/2B fiber diameter between 2 and 6 months of age,
which was partially reversed between 6 and 12 months. The
time course for the reduction in fiber hypertrophy appears to
correlate with the onset of muscle weakness in these animals.
We have generated the first mouse model for human nemaline
myopathy by expressing a dominant negative mutation of
αTmslow. Analyses of these mice clearly demonstrate that the
α-Tmslow(Met9Arg) mutation, identified in a large kindred
with nemaline myopathy, is sufficient for all of the classical
pathologies associated with this disease including rod formation, a
higher proportion of slow, oxidative fibers and muscle
weakness. We have observed that the level of expressed
protein is not predictive of the percentage of rod-containing
fibers in muscles and is not related to muscle usage. A similar
finding was reported for an infant (33) and our results suggest
that this may be a common feature of the disorder. A progressive
hypertrophy occurs in type 2X and 2B fibers in the mouse.
Muscle weakness is apparent when an age-related reversal of
this hypertrophy takes place commencing at 7 months of age.
A compensatory hypertrophy has been reported in similar fiber
types in humans and is most pronounced in those individuals
who display greater muscle strength (15). The mouse model
will serve as a valuable tool to determine the mechanism for
this hypertrophic capability.
Ultrastructural studies of αTmslow(Met9Arg) muscles
showed the presence of two inclusions not normally associated
with nemaline myopathy in humans: cytoplasmic bodies and
tubular aggregates. Both features were observed in a single
muscle type, the SG. Cytoplasmic bodies have been reported
once in association with nemaline myopathy (34) and may
often go unnoticed due to the muscle specificity of the feature.
These features appear to be composed of filamentous material
Human Molecular Genetics, 2001, Vol. 10, No. 4
gene expressed in cultured cardiac myocytes demonstrated
incorporation of the mutant protein into sarcomeres (36). A
requirement of a threshold level of gene expression for the
formation of rods, rather than a graded response, suggests that
one of the factors that contributes to rod formation could be a
loss of integrity of the thin filament. A certain level of incorporated mutant protein may lead to failure of the thin filament
when it is under tension, followed by Z-line streaming and rod
formation. This result may also have implications for a gene
therapy strategy. It is possible that the normal gene under the
control of a suitably strong promoter such as the skeletal actin
promoter may be sufficient to dilute out the deleterious effects
of the mutant protein.
Muscles differ in the extent of rod pathology and hence in
response to the presence of the mutated protein. This cannot be
attributed to differences in fiber type composition since
muscles with similar fiber type composition and similar levels
of protein expression have significantly varied pathology. As
an example, the EDL and SG, both lower hind limb muscles
with comparable fiber type composition, contained 36 and 2%
rod-containing fibers, respectively. It is possible that the intensity
of work done by the muscle has a direct influence on the
muscle pathology. In mouse models for dystrophies such as the
mdx mouse (37) and the desmin knockout mouse (38), a
differential response of the diaphragm, a constantly solicited
muscle, was observed. Indeed, a higher number of rodcontaining fibers in the diaphragm of the nemaline mouse
suggests a basis for nocturnal hypoxia observed in some cases
of nemaline myopathy. Although muscle usage could influence
the number of rod-containing fibers in some muscles in nemaline myopathy, this cannot be the basis for the difference in all
muscles. Shafiq et al. (33) reported 5–15 and 90% rodcontaining fibers in the right and left deltoids, respectively, of
a 10-month-old child with the severe congenital form. Our
finding in the nemaline mice together with this case history has
important implications for the selection of the correct muscle
for human biopsy situations. The diagnosis of nemaline myopathy
may be missed if a biopsy is performed on a muscle with
minimal involvement. More information is required on which
muscles are more susceptible in humans and the factors that
lead to differential pathology in similar muscles.
Figure 7. Fiber type composition of the EDL during maturation. Myosin heavy
chain isoforms (2B, 2A, 1) were detected in frozen sections (20 µm) from EDL
muscles of HSA–αTmslow(Met9Arg) lines 4 and 14 at the ages indicated using
isoform-specific antibodies. Fibers lacking these isoforms were scored as 2X
fibers. Sections from the widest portion of the EDL, containing similar
numbers of fibers, were analyzed. Percentages of fibers expressing a particular
isoform were scored in three to five control (white) and transgenic (black)
animals from one EDL per time point (± SEM).
and may represent a failure in the integrity of thin filaments
due to the mutation in tropomyosin. Tubular aggregates have
not been reported to date in association with human nemaline
myopathy, but have been reported in myopathies associated
with muscle cramps and periodic paralysis (29,30). We and
others (35) have observed tubular aggregates in normal mouse
and rat muscle from 12 months of age. The precocious appearance
of this inclusion in αTmslow(Met9Arg) mice could reflect a
premature aging of the muscle.
Only transgenic lines with an expression level approximately equal to that of the endogenous gene developed rod
pathology. This result demonstrates that a threshold level of
mutant gene expression is required for development of the
disease. It also suggests that the mutant and the wild-type
proteins are competing for the same function in vivo.
Studies using an analogous mutation in the TPM1 (or αTmfast)
Table 1. Contractile parameters of single fibers from EDL of HSA–αTmslow(Met9Arg) line 4 and HSA–αTmslow(wt) line 41 mice
Mean valuesa
No. of fibers
nCa
pCa 50
pCa10
nSr
pSr50
pSr10
Population 1, nemaline
7
1.67
6.23
6.81
5.81
4.56
4.87
0.25
0.14
0.10
1.61
0.05
0.34
5.81
0.12
1.58
6.17
6.78
5.37
4.54
4.72
10.80
1.63
0.27
0.21
0.15
0.78
0.04
0.04
5.76
0.19
2.24
5.68
6.12
6.32c
4.54
4.62c
2.67
1.14
0.48
0.34
0.43
0.82
0.28
0.06
1.82
0.11
2.16
5.76
6.21
5.38
4.54
4.72
4.04
1.22
0.54
0.14
0.24
1.06
0.06
0.07
2.18
0.11
± SD
Population 1, control
8
± SD
Population 2, nemaline
12
± SD
Population 2, control
± SD
11
Force/CSAb
8.72
pCa 50-pSr50
1.67
values ± SD for the major Hill equation parameters calculated for four populations of fibers, two HSA–αTm slow(wt) (control)
and two HSA–αTmslow(Met9Arg) (nemaline) at 9 months of age.
bForce per cross-sectional area in N/cm2.
cSignificance at P < 0.05.
aMean
323
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Human Molecular Genetics, 2001, Vol. 10, No. 4
Figure 8. Muscle strength of HSA–αTmslow(Met9Arg) line 4 in relation to age.
Strength was determined as the percentage pass rate over 15 climbing trials (as
described in Materials and Methods). A lower pass rate over the 15 trials is
indicative of muscle weakness. Pass rates were determined for control (white
bars) and transgenic (black bars) mice at 0–2, 3–4, 5–6, 7–9 and 10–13 months
of age (± SEM). The pass rates were: age 0–2 months, control 87 ± 5% (n =
29) and transgenic 86 ± 5% (n = 20); 3–4 months, control 83 ± 5% (n = 34) and
transgenic 86 ± 4% (n = 37); 5–6 months, control 72 ± 8% (n = 11) and transgenic 55 ± 10% (n = 14); 7–9 months, control 76 ± 8% (n = 13) and transgenic
66 ± 10% (n = 7); 10–13 months, control 66 ± 8% (n = 12) and transgenic
37 ± 7% (n = 24).
αTmslow(Met9Arg) mice have a higher percentage of slow,
oxidative fiber types than non-transgenic littermates. Type 1
fiber predominance is frequently reported in nemaline myopathy;
however, the mechanism for the development of this pathology
has not been resolved. A predominance of type I fibers can
result from a defect in muscle development (3), a conversion
from type 2 to type 1 fibers (2) or a combination of these
mechanisms. The increase in type 1 fibers may represent an
adaptive response to the disease such as that observed in the
mdx mouse diaphragm (39). The mdx mouse diaphragm
undergoes a constant process of degeneration and regeneration
(37) which may facilitate the alteration in fiber type composition.
However, muscle regeneration is not a feature of nemaline
myopathy and, therefore, cannot explain type 1 fiber predominance. From 1 to 12 months of age fiber type composition in
the EDL of HSA–αTmslow(Met9Arg) lines remained relatively
static. This suggests an alteration or defect in the maturation of
fibers as defined by the appearance of the mature myosin
heavy chain isoforms. The percentage of slow/fast, oxidative
fibers was less than that typically seen in human nemaline
myopathy cases. This may reflect the differences in body mass
between the human and mouse and raises the issue of whether
the human myopathic fiber type composition can be achieved
in the mouse.
The responses of individual fibers to Ca2+ and Sr2+ were
examined in skinned fiber experiments. The pCa50 and pSr50
values represent the pCa and pSr corresponding to 50% of ionactivated force, respectively, which give an indication of the
sensitivity of the contractile apparatus to these ions. The values
pCa10 and pSr10 represent the pCa and pSr corresponding to
10% of maximum ion-activated force, and provide a measure
of the threshold of activation. The Hill co-efficient, n, provides
an indication of the minimum number of ions required to activate
one contractile unit of the contractile apparatus and of the level
of co-operativity between the troponin–tropomyosin protein
systems (40). Two different populations of fibers were identified on
the basis of function alone (31,41). The pCa50–pSr50 values of
Figure 9. Hypertrophy in 2X/2B fibers of HSA–αTmslow(Met9Arg) lines.
Fiber diameters were measured from frozen sections of EDL muscles of HSA –
αTmslow(Met9Arg) lines 4 and 14 at (A) 2, (B) 6 and (C) 12 months of age.
Diameters were recorded as the shortest axis across the fiber to the nearest
10 µm. Frequency of occurrence of each fiber diameter is plotted as the mean
percentage of total fibers from two to three control (white bars) and two to four
transgenic (black bars) individuals per time point ( ± SEM).
population 1 are characteristic of 2B fibers and population
2 values are characteristic of 2A fibers (31). Within population 2
fibers, small changes in the nSr and pSr10 values were
observed. There is some evidence to indicate that altering the
isotype of troponin C will influence the force–pSr relationship
(42), thus it is tempting to speculate that there are differences
in force regulation being detected in the population 2 fibers
that can be attributed to the αTmslow(Met9Arg) mutation.
Examination of the normalized force revealed some apparent
differences between the control and nemaline fibers. No
changes in the Ca2+ parameters were observed. The lack of a
change in the pCa50 parameter differs from the finding of
Michele et al. (36), who observed a decrease in this parameter
in a cardiac cell culture model system for this myopathy.
In general, muscle defects in mice result in less severe clinical
phenotypes. This may be due to differences in body mass
between humans and rodents as well as bipedal versus
quadrupedal movement. For example, the mdx mouse model
of Duchenne muscular dystrophy displays relatively mild
muscle pathology and weakness in comparison with humans
Human Molecular Genetics, 2001, Vol. 10, No. 4
(43). Mild muscle weakness has also been observed in the 2B
myosin heavy chain knockout (44), the mouse model for
Bethlem myopathy (45) and a mouse model for myotonic
dystrophy (46). In this respect, the observation of a clinical
phenotype in the nemaline mice is surprising since, in humans,
the αTmslow(Met9Arg) mutation gives rise to a relatively mild
distal myopathy with variable penetrance (47).
The mice develop a late-onset muscle weakness. Weakness
in humans does not correlate with the numbers of fibers
affected with rods or with the degree of type 1 fiber predominance. In the mouse, we were also unable to detect an
unambiguous correlation between these parameters and
muscle weakness. The hypertrophy of fast, glycolytic fibers
that we observed in these mice may be sufficient to prevent
overt muscle weakness in an unchallenged environment.
Hypertrophy of this fiber population has been observed in
some cases of human nemaline myopathy in which muscle
weakness is relatively mild (15). This feature makes the
nemaline mouse a particularly powerful tool to investigate this
innate therapeutic mechanism. The nemaline model described
phenocopies the human condition to a large extent and will be
useful to test potential therapies.
MATERIALS AND METHODS
Transgenic constructs
A 121 base synthetic oligonucleotide carrying the codon
9 ATG→AGG mutation and a synthetic EcoRI site 46 bp
upstream of the translation start site was amplified by PCR and
ligated into EcoRI–BsmI-digested wild-type αTmslow cDNA
(provided by Dr N. Laing). The oligonucleotides used for the
PCR included the forward primer, 5′-TTGCCGAATTCCCAGTTC-3′, the reverse primer, 5′-ATCCAGAGCATTCTCCTTG-3′, and the 121 base oligonucleotide sequence, 5′-TTGCCGAATTCTCCAGTTCTCCAGTGTTCACAGGTGAGCCTACCAACAGCCACTGCTCATGATGGAGGCCATCAAGAAAAAGAGGCAGATGCTGAAGTTAAGACAAGGAGAATGCTCTGGAT-3′ (the mutated base is underlined). The
full-length cDNA carrying the mutation was removed by
EcoRI–PstI digest, ligated into pBlueScript KSII and the mutation confirmed by sequencing. HSA–αTmslow(Met9Arg) was
made by ligating a 1022 bp SmaI–BamHI cassette containing
the intron and 3′-untranslated region (3′-UTR) of SV40 small t
antigen (48) directly downstream of the αTmslow mutant cDNA
sequence. This was followed by blunt-end ligation of a 2.2 kb
HindIII–HindIII fragment of the HSA promoter (48) into the
EcoRV site directly upstream of the mutant αTmslow sequence.
HAS–αTmslow(wt) was created using the same method as for
the previous construct except that the wild-type αTmslow
sequence was inserted. TnIslow–αTmslow(Met9Arg) was made
by blunt-end ligating a 4.2 kb HindIII–HindIII fragment of the
troponin Islow promoter (25) into the EcoRV site. Transgene
constructs were linearized and isolated free of vector
sequences using ClaI–NotI digestion. All enzymes were
supplied by Roche.
Generation and screening of transgenic lines
Fertilized eggs were collected from superovulated FVB/NJ
females on the day of mating, injected with linearized fragment
325
and transferred to pseudopregnant ARC/S females on the same
day according to standard protocols (49). Transgenic mice
from all lines were screened by Southern blot analyses of DNA
extracted from mouse tails (50) using an SV40 3′-UTR probe.
Electron microscopy
Muscles were removed and fixed in 5% glutaraldehyde in
phosphate-buffered saline (PBS) for 4 h at room temperature,
rinsed in PBS and stored in PBS/azide at 4°C. Trimmed
sections (1–2 mm thick) of fixed muscle were post-fixed in 2%
osmium tetroxide (ProSciTech) for 1 h at room temperature,
rinsed in PBS and water, then en-bloc stained with saturated
uranyl acetate for 1 h at room temperature. The specimens
were dehydrated in an ascending series of 50–100% ethanol
and epoxypropane (ProSciTech) and embedded in procure 812
(ProSciTech). Blocks were cut on an ultracut E (Reichart
Young). Sections (70 nm) were cut with a diamond knife
(DDK), stained with uranyl acetate (ProSciTech) and
Reynolds lead citrate (ProSciTech) and examined using a
Phillips EM 400 transmission electron microscope.
Confocal microscopy
Plantaris muscles from 6-month-old HSA–αTmslow(Met9Arg)
line 4 mice were fixed in 2% paraformaldehyde in PBS at room
temperature, transferred to 153 mM ethanolamine for 10 min,
rinsed in PBS and stored in PBS/azide at 4°C until use. Tissues
were embedded in 5% agarose type VII (Sigma) and 100 µm
longitudinal sections obtained with a vibraslice at room
temperature. The sections were blocked with 10% donkey
serum in Tris-buffered saline, 0.1% Tween-20 for 1 h then
labeled with a mouse monoclonal antibody to α-actinin 2
(4B2; 1:500 dilution) (51) for 16 h and visualized with donkey
anti-mouse IgG antibody conjugated to FITC (Jackson Laboratories). Sections were mounted with Vectashield (Vector
Laboratories) and 1 µm optical sections were obtained with a
confocal microscope (Leica).
Northern blot analysis
Animals were sacrificed at 2 months of age by cervical
dislocation and muscles were snap frozen in liquid nitrogen
and stored at –80°C prior to use. Total RNA was extracted
from muscle by the TRIzol method (Life Technologies). Quantitative northern blot analysis was performed on 10 µg of total
RNA from either the crural muscle block (excluding soleus)
from HSA–αTmslow(Met9Arg) lines or soleus from TnIslow–
αTmslow(Met9Arg) lines according to the protocol of
Sambrook et al. (50). An oligonucleotide probe to the first 180
bases in the coding region of the αTmslow gene was hybridized
(52) and washed at 65°C using established protocols (50). The
probe recognizes only the 1.3 kb endogenous αTmslow and the
2.4 kb transgene transcripts. Loading errors were corrected by
hybridizing the blots with an 18S rRNA-specific oligonucleotide probe under conditions of probe excess. Levels of transcripts were quantified by densitometric analysis (ImageQuant
v4.0; Molecular Dynamics) of images obtained from a
STORM 860 phosphorimager (Molecular Dynamics). HSA–
αTmslow(Met9Arg) and HSA–αTmslow(wt) transcript levels
were expressed as a percentage of the endogenous αTmslow
transcripts from type 1 fibers in non-transgenic soleus (55%
326
Human Molecular Genetics, 2001, Vol. 10, No. 4
type 1) (53). TnIslow–αTmslow(Met9Arg) transcript levels were
expressed as a percentage of the endogenous αTmslow transcripts in the same muscle.
Protein extracts and western blotting
Mice were sacrificed at 2 months of age and the EDL, SG, TA,
plantaris, soleus, diaphragm, flexor carpi ulnaris, flexor carpi
radialis, flexor digitorum profundus and ECU muscles were
excised, snap frozen in liquid nitrogen and stored at –80°C
prior to use. Crude myofibrillar preparations were made from
frozen muscle samples, crushed with a mortar and suspended
in 10 mM Tris pH 7.4, 5 mM MgCl2. Samples were treated
with 2.5 µg of RNase A (Roche), sonicated on ice (20 × 1 s
pulses), followed by the addition of 50 U of DNase I (Roche)
and incubation on ice for 15 min. Following centrifugation at
10 000 g, 4°C for 10 min, the pellet was resuspended in 8 M
urea, 40 mM dithiothreitol, 0.5% Triton X-100 and 2.5%
Ampholine pH 3–10 (Sigma). Protein concentration was
calculated by Bradford assay (Bio-Rad) according to standard
protocols (54). Isoelectric focusing was performed on
immobiline dryplate pH 4–7 gels (Amersham Pharmacia
Biotech) prepared according to the manufacturer’s instructions
and the protocol described by Holmquist (55). Fifteen
micrograms of skeletal muscle extract was loaded for each
sample and focused at 300 V, 5 mA for 1 h then 2000 V, 5 mA
for 4 h. Gels were transferred to nitrocellulose membrane (Bio-Rad)
and incubated with a mouse monoclonal anti-sarcomeric Tm
antibody CH1 (Sigma) followed by a secondary rabbit
antimouse IgG antibody conjugated to horseradish peroxidase
(HRP) (Bio-Rad). Bands were visualized by treating with
Lumi-lightPLUS western blotting substrate (Roche) and
exposing to autoradiography film. Loading was assessed to be
equivalent by immunoblot using an antibody (C4; kindly
supplied by Dr Jim Lessard) which recognizes all cytoplasmic
and sarcomeric actins (56). Autoradiographs were digitized
using a Molecular Dynamics computing densitometer and
analyzed using ImageQuant v4.0 software (Molecular
Dynamics).
Quantification of nemaline rods
Rod quantification was performed on muscles collected and
prepared as described for electron microscopy. Sections
(0.3 µm) were stained with toluidine blue for light microscopy.
Rods were identified by morphology and by similar staining
intensity with the Z-line. The numbers of fibers affected with
rods were calculated from six sections of three animals per
time point. Rod dimensions were measured from electron
micrographs, calibrated using a cross-grating replica. At least
85 rods were measured per muscle. Statistical analysis of data
was performed using single-factor analysis of variance
(ANOVA) (MS Excel 2000).
Quantification of fiber type and diameter
EDL muscles from 1-, 2-, 6- and 12-month-old HSA–
αTmslow(Met9Arg) mice, lines 4 and 14, were removed,
stretched to prevent contracture, coated in tissue-freezing
medium (ProSciTech), frozen in isopentane cooled with liquid
nitrogen and stored in liquid nitrogen. Samples were
equilibrated to –24°C prior to sectioning. Cryostat sections
(20 µm) taken from the midsection of the EDLs were fixed in
2% paraformaldehyde in PBS on ice for 2 min, rinsed in PBS
and blocked with 10% normal goat serum in PBS for 30 min.
Consecutive sections were treated with the following antibodies to detect isoforms of myosin heavy chain (MyHC):
culture supernatant from hybridomas secreting antibody to
type 1 (BA-F8; undiluted) (57), 2A (SC-71; undiluted), 2B
(BF-F3; 1:10 dilution) (58), or 2A + 2B (NCL-MHCf;
1:50 dilution) (Novacastra), and purified antibody that
recognizes all adult isoforms but 2X (BF-35; 1:200 dilution)
(58). All antibody dilutions were made with 2% normal goat
serum in PBS. BA-F8, SC-71, BF-F3 and BF-35 hybridomas
were purchased from the German Collection of Microorganisms
and Cell Cultures. MyHC-positive fibers were visualized using
immunoperoxidase detection as described by Schiaffino et al.
(58) using a goat-anti-mouse secondary antibody (Bio-Rad)
conjugated to HRP (1:200 dilution) and a DAB substrate kit
(Vector Laboratories). Fiber diameter was determined by
measuring the shortest axis for all fibers within a section to the
nearest 10 µm. Sections were analyzed from two gendermatched individuals at each time point.
Skinned fiber experiments
Dissections of EDL from HSA–αTmslow(Met9Arg) line 4 mice
and HSA–αTmslow(wt) line 41 mice were carried out under
paraffin oil at 4°C, following which the fibers were chemically
‘skinned’ using a mixture of a relaxing solution and 2% Triton
detergent. The muscle fibers were activated by a series of solutions
containing calcium and strontium ions in buffered concentrations, following the formulae of Fink et al. (31). Solutions of
differing concentrations (measured as pCa = –Log10[Ca2+] and
pSr = –Log10[Sr2+]) were obtained by mixing a relaxing solution
containing EGTA2– (50 mM) in various proportions with a
maximally activating solution containing either Ca2+ or Sr2+.
Experiments were carried out using the methods of Stephenson
and Williams (41). Skinned fiber segments (typically 500–
1000 µm long) were mounted between a pair of sharpened
jeweler’s forceps and a stainless steel pin using fine surgical
silk suture (Deknatel 10/0). The pin was attached to a piezoresistive strain gauge (type AME301), the output of which was
amplified and displayed oscillographically. Sarcomere length
was adjusted to between 2.60 and 2.80 µm using a He–Ne laser
diffraction method as well as by direct microscopic observation.
Statistical analysis was undertaken by the use of Student’s
t-test between the means of each curve parameter. Significance
was flagged if occurring at less than the 0.05 level.
Strength and fatigability test
Whole animal strength and fatigability was carried out
according to the method of Hübner et al. (32). Animals were
placed with their forepaws on a metal rod covered in heat
shrink rubber with a diameter of 3 mm. The mice were required
to pull themselves up on top of the rod to pass the test. This was
repeated 15 times within 3 min. Mice that were unable to pull
themselves on top of the rod or fell were classed as a fail. The
measurement of muscle weakness was based on the mean
percentage of passes over the 15 trials. Animals were pooled into
five age groups. The number of HSA–αTmslow(Met9Arg) line
4 and non-transgenic littermates assessed are shown in parentheses: 0–2 months (29, 20), 3–4 months (34, 37), 5–6 months
Human Molecular Genetics, 2001, Vol. 10, No. 4
(11, 14), 7–9 months (13, 7), 10–13 months (12, 24). Statistical
anlaysis of each time point was performed by t-test assuming
unequal variances (MS Excel 2000).
ACKNOWLEDGEMENTS
We wish to thank Carina Wallgren-Pettersson, Nigel Laing,
Victor Dubowitz and members of the International Consortium
on Nemaline Myopathy, R. Weinberger, P. Robinson and C.
Vockler for valuable advice and discussions; V. Hodgson, S.
Kim, E. Miles and J. Cuomo for technical assistance; Stefano
Schiaffino for purified BF-35 antibody; Prof. P. Rowe for
supporting this work. We are indebted to Luana Ferrara,
Wendy Thornton and staff of the CMRI animal facility. This
work was supported by grants from the Royal Alexandra
Hospital for Children to E.C.H. and K.N.N. and from the
National Health and Medical Research Council of Australia to
E.C.H., G.F.D. and K.N.N.
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