Human Molecular Genetics, 2001, Vol. 10, No. 4 317-328
© 2001 Oxford University Press
A mutation in 
-tropomyosinslow affects muscle strength, maturation and hypertrophy in a mouse model for nemaline myopathy
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.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Nemaline myopathy is a hereditary disease of skeletal muscle defined by a distinct pathology of electron-dense 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
-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 |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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, adult-onset 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 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 -actinin (4,5). Ultrastructural studies also have suggested that sarcomeric -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 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: Tmslow (16,17), nebulin (18) and -skeletal actin (19). -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 disease-associated 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 Tmslow (TPM3), likely resulting in a dominant negative mutation. A second mutation with recessive inheritance has also been identified in Tmslow 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 and ß subunits (22). The tropomyosin filament lies in the major groove of filamentous -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 -actinin and actin resulting in the extension of -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 Tmslow(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 rod-containing 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 hypertrophy that correlates with the onset of muscle weakness which occurs at 5–6 months of age.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Generation of mice expressing the
Tmslow(Met9Arg) transgeneWe 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 fiber-specific expression of the mutant protein. In order to achieve this, we linked the coding region for
Tmslow(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–Tmslow(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 Tmslow(Met9Arg) in type 2 fibers of the mouse. In this construct, the coding region of Tmslow(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–Tmslow(Met9Arg) (Fig. 1B). To establish that any pathological features arising in the HSA–Tmslow(Met9Arg) 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 Tmslow protein was linked to the HSA promoter region. Nine founder lines were generated with this construct designated HSA–Tmslow(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–Tmslow(Met9Arg), HSA–Tmslow(Met9Arg) and HSA–Tmslow(wt) lines of mice unless otherwise indicated.
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Mice expressing
Tmslow(Met9Arg) develop nemaline rodsNemaline rods were present in the slow fiber-containing soleus and complex zone of the crural muscle block in one transgenic line expressing the TnIslow–
Tmslow(Met9Arg) construct and in all fast fiber-containing muscles of three lines expressing the HSA–Tmslow(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–Tmslow(wt) which established that inappropriate expression of Tmslow 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 -actinin 2 (Fig. 3), a major component of nemaline rods and the Z-line (4,5). This result unambiguously demonstrates that the Tmslow(Met9Arg) mutation is responsible for rod pathology in humans with this TPM3 mutation and that it acts through a dominant mechanism (17).
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Mice expressing
Tmslow(Met9Arg) develop cytoplasmic bodies and tubular aggregates of the sarcoplasmic reticulumTwo inclusions not typically associated with nemaline myopathy in humans were observed only in the HSA–
Tmslow(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).
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Rod formation correlates with a threshold level of
Tmslow(Met9Arg) mRNARod pathology was observed in those lines of mice in which a level of
Tmslow(Met9Arg) mRNA approximating that of the endogenous Tmslow was achieved. Tmslow(Met9Arg) transcript levels were determined in the crural muscle block (without soleus) of HSA–Tmslow(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 Tmslow gene in slow fibers (Fig. 5). Further support for this threshold phenomenon was observed in the highest expressing TnIslow–Tmslow(Met9Arg) line 49. When the transgenic locus in this line is in the hemizygous state, the Tmslow(Met9Arg) transcript level is 50% of the endogenous Tmslow 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 Tmslow(Met9Arg) locus must approach levels comparable to the endogenous Tm loci in order for rods to form. This is analogous to the mutant to normal Tmslow ratio found in the human condition.
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The number of fibers containing rods and rod size are different in different muscles
The HSA–
Tmslow(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 rod-containing 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. -tropomyosinfast (Tmfast) and ß-tropomyosin (ßTm) run as a doublet as the lower (most acidic) band. In slow muscle (soleus) extracts, the band running below the mutant and above the Tmfast and ßTm bands is the Tmslow 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–Tmslow(Met9Arg) line 49 mice since only two muscles in this line of mice contain rods.
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) 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–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 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–Tmslow(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–Tmslow(Met9Arg) lines remained relatively static, suggesting a disruption to normal maturation. A parallel analysis of HSA–Tmslow(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–Tmslow(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–Tmslow(Met9Arg) line 49 mice was similar to wild-type. 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.
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Skinned fibers from nemaline mice show minor changes in response to Sr2+
Single fiber mechanical studies were performed on 15 fibers from HSA–
Tmslow(Met9Arg) line 4 mice and 23 fibers from HSA–Tmslow(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.
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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–
Tmslow(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–Tmslow(Met9Arg) line 4 mice in comparison with nontransgenic littermates. At 5–6 months of age, HSA–Tmslow(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 non-transgenic littermates (P < 0.05). The late onset of muscle weakness is reminiscent of the childhood-onset disorder observed in humans with the Tmslow(Met9Arg) mutation.
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Mice expressing
Tmslow(Met9Arg) display hypertrophy of fast, glycolytic fibersIn 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–
Tmslow(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–Tmslow(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/2B fibers is greater in HSA–Tmslow(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. 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.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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We have generated the first mouse model for human nemaline myopathy by expressing a dominant negative mutation of
Tmslow. Analyses of these mice clearly demonstrate that the -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 Tmslow(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 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 Tmslow(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 Tmfast) 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 rod-containing 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% rod-containing 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.
Tmslow(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–Tmslow(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 ion-activated 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 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 Tmslow(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 (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 Tmslow(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.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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 Tmslow 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–Tmslow(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 Tmslow 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 Tmslow sequence. HAS–Tmslow(wt) was created using the same method as for the previous construct except that the wild-type Tmslow sequence was inserted. TnIslow–Tmslow(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 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–Tmslow(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 -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–Tmslow(Met9Arg) lines or soleus from TnIslow–Tmslow(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 Tmslow gene was hybridized (52) and washed at 65°C using established protocols (50). The probe recognizes only the 1.3 kb endogenous Tmslow 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–Tmslow(Met9Arg) and HSA–Tmslow(wt) transcript levels were expressed as a percentage of the endogenous Tmslow transcripts from type 1 fibers in non-transgenic soleus (55% type 1) (53). TnIslow–Tmslow(Met9Arg) transcript levels were expressed as a percentage of the endogenous Tmslow 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 x 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–Tmslow(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 gender-matched individuals at each time point.
Skinned fiber experiments
Dissections of EDL from HSA–Tmslow(Met9Arg) line 4 mice and HSA–Tmslow(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 piezo-resistive 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–Tmslow(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 (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).
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ACKNOWLEDGEMENTS |
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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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FOOTNOTES |
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+ To whom correspondence should be addressed. Tel: +61 2 9687 2800; Fax: +61 2 9687 2120; Email: ehardeman@cmri.usyd.edu.au
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REFERENCES |
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