Human Molecular Genetics, 2000, Vol. 9, No. 9 1393-1402
© 2000 Oxford University Press
Myopathy phenotype of transgenic mice expressing active site-mutated inactive p94 skeletal muscle-specific calpain, the gene product responsible for limb girdle muscular dystrophy type 2A
1Department of Molecular Biology, Institute of Molecular and Cellular Biosciences, University of Tokyo, Tokyo, Japan, 2Department of Physiological Chemistry and Metabolism, Graduate School of Medicine, University of Tokyo, Tokyo, Japan, 3National Institute of Neuroscience, NCNP, Tokyo, Japan, 4Department of Laboratory Animal Science, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan, 5Department of Immunology, Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan and 6Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, Tokyo, Japan
Received 25 January 2000; Revised and Accepted 17 March 2000.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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A defect of the gene for p94 (calpain 3), a skeletal muscle-specific calpain, is responsible for limb girdle muscular dystrophy type 2A (LGMD2A), or ‘calpainopathy’, which is an autosomal recessive and progressive neuromuscular disorder. To study the relationships between the physiological functions of p94 and the etiology of LGMD2A, we created transgenic mice that express an inactive mutant of p94, in which the active site Cys129 is replaced by Ser (p94:C129S). Three lines of transgenic mice expressing p94:C129S mRNA at various levels showed significantly decreased grip strength. Sections of soleus and extensor digitorum longus (EDL) muscles of the aged transgenic mice showed increased numbers of lobulated and split fibers, respectively, which are often observed in limb girdle muscular dystrophy muscles. Centrally placed nuclei were also frequently found in the EDL muscle of the transgenic mice, whereas wild-type mice of the same age had almost none. There was more p94 protein produced in aged transgenic mice muscles and it showed significantly less autolytic degradation activity than that of wild-type mice. Although no necrotic–regenerative fibers were observed, the age and p94:C129S expression dependence of the phenotypes strongly suggest that accumulation of p94:C129S protein causes these myopathy phenotypes. The p94:C129S transgenic mice could provide us with crucial information on the molecular mechanism of LGMD2A.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Calpain (EC 3.4.22.17) is an intracellular cysteine protease, which is regulated by Ca2+ (1–4). The conventional mammalian calpains, µ- and m-calpain, are ubiquitously expressed, suggesting their involvement in basic and essential cellular functions mediated by the Ca2+ signaling pathway. p94 (also called calpain 3, nCL-1 or CAPN3) is a skeletal muscle-specific calpain homolog that shows overall similarity to the conventional calpain catalytic subunit (5,6). In addition to its predominant expression in skeletal muscle, p94 has several unique properties. It shows highly rapid and exhaustive autolytic activity. Its in vitro half-life is <10 min (7,8). In the course of researching p94 stabilizing factor in vivo, we found that p94 specifically binds to connectin (also called titin), a gigantic filamentous muscle protein (9). The molecular mass of connectin is >3 x 106 and it is ~1 µm long, connecting the skeletal muscle Z- and M-lines by a single molecule (10–13).
p94 protein binds to the N2A region of connectin around one of the p94-specific sequences, IS2, which is also involved in rapid autolysis (7). Connectin tends to split at the N2 line, which is located close to the N2A region, and the conventional calpains are responsible for this proteolysis (14,15). In contrast, p94 does not cut connectin, at least in this region. Although the physiological significance of binding between p94 and connectin is still unclear, these findings lead to the idea that binding to the N2A region of connectin stabilizes p94 and that without this binding p94 rapidly disappears. In other words, it is possible that connectin serves as a ‘base’ for p94 to function.
Recently, a defect in the p94 gene was shown to be responsible for limb girdle muscular dystrophy type 2A (LGMD2A), or ‘calpainopathy’ (16,17). LGMD2A is characterized by progressive muscle degeneration in the pelvic and shoulder girdles, showing autosomal recessive inheritance with considerable prevalence (18–20). In patients with LGMD2A, more than 100 independent pathogenic mutations in the p94 gene have been reported, including more than 60 point mutations (17,20–22). LGMD2A is unique in that a defect of a gene encoding an enzyme causes muscular dystrophy. Other muscular dystrophies are caused by deficiencies of structural proteins around the sarcolemma or at the nuclear inner membrane (23–26). Our recent study has revealed that several missense point mutations of p94 found in LGMD2A commonly lead to a deficiency of proper proteolytic activity on substrates such as fodrin (27). This indicates that the molecular mechanism for LGMD2A is distinct from that for the other muscular dystrophies.
To clarify the physiological functions of p94 in relation to connectin and to elucidate the molecular mechanism of LGMD2A, we have constructed transgenic mice that express the active site Cys129
Ser mutant of p94 (p94:C129S). It is largely believed that the CysSer mutation does not alter molecular conformation, as several enzymes that have Ser in the active site have been shown to retain significant activity, even if the Ser is changed to Cys or vice versa (28–30). Our previous experiments revealed that Cys129 is the most important residue for proteolytic activity of p94 and that the C129S mutation of p94 completely stopped the rapid autolysis referred to above (7). In other words, p94:C129S protein is expressed stably in cultured cells, whereas the wild-type protein is very unstable and hardly detectable. Thus, regardless of the recessive nature of LGMD2A, we hypothesized that transgenic expression of p94:C129S might show a dominant-negative effect on wild-type p94 and thus we created and examined the requisite transgenic mice.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Generation of transgenic mice expressing p94:C129S, a protease-inactive mutant form of p94
From 99 transgene-injected eggs, 95 mice were born. Among them, 90 mice grew normally, resulting in 19 founders that possessed the p94:C129S transgene, as described in Materials and Methods. The appearance and behavior of the p94:C129S transgenic founder mice did not apparently differ from those of wild-type mice. Among 167 mice in the F1 and F2 generations produced by mating the founder mice with wild-type mice, 78 had the p94:C129S transgene (38 females and 40 males). These data suggest that p94:C129S expression did not result in lethal effects, nor did it affect the sex ratio. The female transgenic mice, however, became pregnant less frequently than did wild-type mice and the mothers killed their pups more frequently than did wild-type mice. Finally, we have established three lines of p94:C129S transgenic mice (called S21, S44 and S62) from independent founders. As shown in Table 1, no significant difference in body weight was observed between the transgenic and wild-type mice at 8 and 12 weeks of age. However, at 20 and/or 32 weeks, the S44 and S62 lines showed significantly increased weight.
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Detection of p94:C129S protein in the transgenic mice
To confirm that p94:C129S transgenic mice are expressing p94:C129S protein, we analyzed a skeletal muscle extract by immunoblotting methods, using p94-specific anti-IS2 antiserum (27). We detected p94 protein in the crude connectin fraction from skeletal muscle of the transgenic as well as the wild-type mice, as described in Materials and Methods. However, the amount of p94:C129S protein was not clearly determined, as it was impossible to distinguish p94:C129S from endogenous wild-type p94. Total amounts of p94:C129S and wild-type p94 proteins were not different between the 32-week-old transgenic and wild-type mice (Fig. 1). We could not detect p94 protein with anti-IS2 antiserum in extracts of liver, spleen, heart and whole blood cells, suggesting that exogenous p94:C129S protein is degraded in these organs and/or that the expression level of p94:C129S protein is low.
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Therefore, we attempted to quantify the transgene mRNA to distinguish p94:C129S from endogenous wild-type p94. Competitive PCR quantification of p94:C129S mRNA in skeletal muscle cDNA revealed that the amount of p94:C129S mRNA in S62 is ~10% of that of wild-type p94. For S21 and S44, although transgene mRNA was detected, the detection level was below the quantification range, suggesting that the amounts are near or less than 1% of that of wild-type. Next, we examined the relative amounts of p94:C129S mRNA among lines S62, S44 and S21. Using whole blood cells as a source of cDNA, we quantified the amounts of transgene mRNA, normalized to the amounts of ß-actin mRNA. As shown in Figure 2, the average amounts of p94:C129S mRNA in the S62, S44 and S21 lines were 72, 9.7 and 1.4 copies/1000 copies of ß-actin, respectively. In other words, the relative expression levels in whole blood cells were 100:13:1.9, consistent with the above observations in skeletal muscle. Although the absolute level of transgene expression was not very high, we confirmed its expression in skeletal muscle and have started to examine various characteristics of the transgenic mice.
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Analyses of the motor function of the transgenic mice
To investigate whether expression of p94:C129S affects muscle functions, we examined motor function by measuring grip strength. This has been commonly used as an indicator of muscle strength following administration of muscle relaxants or neurotoxic reagents (31). As shown in Figure 3, grip strength showed p94:C129S expression level-dependent and growth-dependent decreases. The grip strength values of 32-week-old female S44 mice and of 20-week-old female S62 mice were statistically distinct from wild-type mice at the 5% level of significance. Moreover, 32-week-old female S62 mice showed depression at the 0.5% level of significance. No significant difference was observed for the S21 mice. These data strongly suggest that expression of p94:C129S protein perturbs muscle motor function in a dose-dependent manner, assuming that whole blood cell mRNA levels reflect those of skeletal muscle. Furthermore, the growth-dependent muscle weakness of the transgenic mice coincides with the pathological feature of limb girdle muscular dystrophy.
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Histochemical analysis of skeletal muscle fibers
We analyzed histochemical features of the transgenic mice, using skeletal muscle sections from the transgenic and wild-type mice. The extensor digitorum longus (EDL) and soleus muscles were stained using hematoxylin and eosin (H&E), modified Gomori trichrome (mGT) and NADH–tetrazolium (NADH-TR) methods. Cryostat frozen sections of 40-week-old S62 mice showed no significant differences from those of wild-type (Fig. 4). Similarly, muscle fibers of 40-week-old S21 and S44 mice also showed no abnormalities (data not shown).
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Since the LGMD2A phenotype becomes more severe upon aging, we suspected that older transgenic mice would show clearer histochemical symptoms. The EDL and soleus muscle fibers of two 106-week-old S62 mice and two 106-week-old wild-type mice were examined by the same staining methods used above. A number of strikingly abnormal aggregates, stained bright red by the mGT method, were found in EDL muscle fibers of 106-week-old S62 mice (Fig. 5b). The frequencies of occurrence were 38 and 22% of the total fibers in two independent S62 mice, whereas two independent wild-type mice showed none in preparations that we observed (Table 2 and Fig. 5e). While they are not specific features of LGMD2A, these aggregates resembled the so-called ‘tubular aggregates’ that are often observed in the skeletal muscles of patients with a variety of different disorders. These include periodic paralysis, related myotonic disorders, malignant hyperthermia, inflammatory myopathy and myopathy after exposure to alcohol and various drugs (32).
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Nuclei of EDL muscle fibers of two independent 106-week-old S62 mice were observed in the central area of the fibers (Figs 5a and 6). The frequencies of such fibers were 7.6 and 6.9%, while the frequency in wild-type mice was <1% (Table 2 and Fig. 5d). In soleus muscles, on the other hand, the nuclei were located at the periphery of the fibers in both S62 and wild-type mice (Fig. 5g and j).
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Lobulated fibers were found in the soleus muscles of two 106-week-old S62 mice (Fig. 5i), at frequencies of 14 and 25% of total fibers (Table 2). In soleus muscles of 106-week-old wild-type mice lobulated fibers were rarely observed (Fig. 5l), with a frequency <10% (Table 2). Accumulation of mitochondria-like structures at the periphery of soleus muscle fibers (Fig. 5g–l) was observed in both 106-week-old S62 and wild-type mice, suggesting that this finding might result from aging. Significant fiber splitting and split fibers in addition to central nuclei were sometimes observed in EDL muscle sections of 106-week-old S62 mice, as shown in Figure 6, whereas wild-type mice showed none (data not shown). No necrotic–regenerative pattern, a central feature of all muscular dystrophies, was observed at any time.
Recently, possible involvement of apoptosis in LGMD2A has been reported (33). The transgenic mice were examined for the presence of apoptosis using the terminal deoxynucleotide transferase-mediated dUTP–biotin nick end labeling (TUNEL) method or immunostaining with activated caspase 3- and caspase 9-specific antibodies (34,35). However, no or almost no positive signals were detected by any of these methods, as in the case of wild-type mice (data not shown).
Increased amount and reduced autolytic degradation activity of p94 protein in the crude connectin fraction of 106-week-old S62 transgenic mouse muscle
To confirm the accumulation of p94:C129S protein in 106-week-old S62 transgenic mice, which showed microscopic abnormalities as mentioned above, we examined the p94 protein of muscles of S62 transgenic and wild-type mice. If the p94:C129S protein accumulates in the aged transgenic mice, the ratio of p94:C129S to endogenous wild-type p94 protein should increase, resulting in a reduction in the autolytic degradation activity of p94 protein as a whole. Total p94 protein contents in the crude connectin fraction from two independent 106-week-old S62 transgenic and wild-type mice were detected by p94-specific antiserum. Initially, there was slightly more p94 protein in 106-week-old S62 transgenic mice, compared with wild-type mice of the same age, as shown in Figure 7a. Densitometry of the bands indicated that the band density of p94 (relative to myosin heavy chain) of the transgenic mice was approximately twice as large as that of the wild-type mice (see Fig. 7a legend). Next, these fractions were incubated at 0 or 37°C for 15 min to determine their autolytic degradation activity. The p94 protein from wild-type mouse muscle showed rapid degradation of the 94 kDa band with an increase in the 55 kDa degradation intermediate fragments, whereas that from the transgenic mice was stable (Fig. 7b). These results indicate that p94:C129S protein supplants the endogenous wild-type p94 protein and accumulates in muscles of S62 transgenic mice, resulting in inhibition of proteolytic activity of endogenous wild-type p94.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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In this study we have established three lines of transgenic mice ubiquitously expressing p94:C129S. The relative mRNA levels of p94:C129S were S62 >> S44
S21, although the protein concentrations were not determined. They did not show any different appearance from that of wild-type mice at the early stages, at least until 12 weeks after birth. After 20 weeks, however, S62 mice weighed more than wild-type. One possible explanation for this result is that S62 mice are not as active as wild-type mice, owing to a slight muscle weakness as measured by grip strength. We observed that S62 mice had much more adipose tissue compared with the wild-type mice when dissected (data not shown). Weakened grip strength is significant for S62 mice, although very mild and almost undetectable for lines S44 and S21, respectively, corresponding to their levels of p94:C129S mRNA expression. However, we cannot eliminate the possibility that the observed difference occurred through side-effects of the transgene, such as disruption of an unrelated gene(s). Histochemical analyses of sections of 106-week-old S62 mice, however, clearly showed microscopic abnormalities, such as centrally placed nuclei, tubular aggregate-like structures and lobulated and split fibers. Such findings were only rarely observed in wild-type mice of the same age. Moreover, the grip strength of S62 mice was significantly reduced compared with that of wild-type and the grip strength decreased as S62 mice grew older. In contrast, muscle fibers of 40-week-old mice did not show any of the above abnormalities.
The amount of p94:C129S mRNA in skeletal muscle of three transgenic mouse lines was between ~10% and less than 1% of endogenous wild-type p94 mRNA. The amount of total p94 proteins (wild-type mouse p94 + rat p94:C129S) detected in skeletal muscle of the 32-week-old transgenic mice was almost identical to that of wild-type mice. These results seem to indicate that p94:C129S protein in the transgenic mice is at a very low level at this stage. The actual ratio of p94:C129S protein to wild-type protein could not be determined, because the p94:C129S protein cannot be distinguished from the endogenous wild-type protein by immunoblotting.
However, 106-week-old S62 mice had increased quantities of p94 protein in skeletal muscle and it showed reduced autolytic degradation activity. These observations can be explained by the following hypothesis. Previously, we reported that the immunochemically detected level of wild-type p94 protein in COS7 cells is <1% of that of p94:C129S, when expressed under the same promoter (7). Thus, the turnover of p94:C129S protein is much slower than that of the wild-type protein. As a result, p94:C129S protein binding to connectin slowly icreases with time and finally predominates over the wild-type p94 protein. On the other hand, cytosolic p94 protein (both wild-type and p94:C129S), dissociated from connectin, should rapidly autolyse or be proteolysed by wild-type p94 and/or other proteases. This leads to a situation where only p94 bound to connectin can exist in a stable form in skeletal muscle. It is likely that the total amount of p94 protein in any one myocyte is regulated at a certain level by connectin binding, since wild-type p94 (the form not binding to connectin) is very unstable in vitro. Moreover, there are constant and limited numbers of binding sites for connectin in the myofibril. Therefore, we conclude that S62 transgenic mice probably exhibit a myopathy phenotype due to interference with p94 protease activity followed by accumulation of p94:C129S protein.
An intriguing aspect of this study is the apparent discrepancy that transgenic mice that also express wild-type p94 showed a dysfunctional phenotype, while LGMD2A is inherited in an autosomal recessive manner. No phenotype should be observed if a mouse has at least one wild-type p94 allele. This point can be explained as follows.
The C129S mutation, which completely abolishes proteolytic activity without altering the protein 3-dimensional conformation, has not yet been found as a LGMD2A allele, although more than 100 distinct pathogenic p94 mutations have been identified (17,20–22). p94:C129S looks like the ‘normal’ protein and may, therefore, escape cellular degradation systems, resulting in accumulation of the protein and domination over the wild-type protein with time. It is also possible that the C129S mutation has different properties compared with other LGMD2A mutations. Other p94 mutants found in LGMD2A patients may be identified as ‘abnormal’ proteins by the cell system so that they are rapidly degraded by, for example, the ubiquitin–proteasome system. Indeed, some LGMD2A mutant p94 is stably expressed in vitro due to loss of rapid autolytic activity (27), whereas it can barely be detected in vivo (36,37). In these heterozygotes, wild-type p94 can function without being disturbed by the mutant p94.
Alternatively, the gene promoter that we used to express p94:C129S mRNA in mice is a ubiquitous viral one. Thus, the spatio-temporal expression pattern may differ from that of the endogenous p94 gene. On the other hand, both alleles of a heterozygote for the p94 gene, as in a LGMD2A carrier, are under the control of the same p94 gene promoter. The following two possibilities can be considered here. (i) At early developmental stages expression of the wild-type p94 gene is very low (38), suggesting that p94:C129S expression may predominate. Thus, it is likely that this perturbation at early stages influences later development of the skeletal muscle. (ii) Even in adult skeletal muscle cells, expression of wild-type p94 varies, depending on muscle cell type, as in the case of conventional calpains (39,40). In this case, although the total amount of p94:C129S mRNA expressed is less than that of wild-type, some cells that express very small amounts of wild-type p94 may predominantly express p94:C129S. Abnormalities may appear only in these cells, resulting in mild dysfunction of the total muscle.
Interestingly, both autosomal dominant myotonia congenita (Thomsen’s disease) and recessive generalized myotonia (Becker’s disease) are caused by different mutations in the same gene encoding the major skeletal muscle Cl– channel (CLCN1, 7q35) (41). Mutations such as G230E cause Thomsen’s disease, whereas R496S, etc., which simply destroy channel function, cause Becker’s disease. Therefore, it is possible that some unidentified p94 mutations may show different modes of inheritance.
From the results in this study, together with the previous findings on the functions of p94, we propose the following insights into the molecular mechanism of LGMD2A.
(i) That expression of p94:C129S causes myopathy phenotypes indicates that a p94 protein defect could be the sole cause of LGMD2A. Moreover, the p94 protein itself, and not the gene, is definitely responsible for LGMD2A symptoms, although other factors or genes may be involved in LGMD2A.
(ii) As is also indicated by our previous studies (27), proteolytic activity of p94 is essential for normal function of skeletal muscle. Although the in vivo proteolytic target(s) of p94 has not yet been determined, we have identified several in vitro substrates for p94 (Y. Ono, H. Sorimachi, S. Tomioka, G. Hashimoto, S. Ishiura and K. Suzuki, manuscript in preparation) that are candidates for in vivo targets. Some of these may provide us with crucial information about LGMD2A.
(iii) As mentioned above, the lack of a significant difference between the total protein levels of p94 in crude connectin fractions of the transgenic and wild-type mice strongly suggests that the amount of p94 binding to connectin is regulated. It is likely that the rapid autolytic activity of p94 is a major part of this mechanism. This result also suggests that unbound p94:C129S, which has no autolytic activity, is also rapidly degraded in skeletal muscle cytosol. This may be accomplished by intermolecular proteolysis, by the wild-type p94 or some unknown p94-degrading protease system. That accumulation of p94:C129S was not observed in young transgenic mice can be partly explained by these proteolytic events. Elucidation of the p94-degrading enzyme system(s) will give us another clue to understand the molecular mechanism of LGMD2A.
It seems odd that inactivation of a protease leads to muscle degeneration including protein degradation. It is believed that in Duchenne type muscular dystrophy and other muscular dystrophies final activation of the conventional calpains is achieved by an increased Ca2+ concentration, caused in turn by a destabilized membrane structure lacking dystrophin, sarcoglycans and/or other muscle structural proteins. Thus, p94 must function in a repressive manner, at least against final protein degradation. A similar case has been found in the Saccharomyces cerevisiae alkaline adaptation process that we reported recently (42). CPL1, which encodes a calpain-like cysteine protease, is involved in activation of the Rim101p transcription factor that is responsible for expression of various enzymes under alkaline conditions. When CPL1 is disrupted, Rim101p also disappears, indicating that Cpl1p protease functions to stabilize Rim101p. One likely speculation is that p94 as well as Cpl1p proteolytically inactivate proteases or an activator of the proteases that are responsible for muscle degeneration. Thus, p94 and Cpl1p represent a novel type of protease that modulates intracellular proteolytic events. The molecular mechanism of LGMD2A must relate to dysfunction of p94 as an intracellular proteolytic system modulator.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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The transgene construction
The expression plasmid pSRD-p94:C129S was reported previously (ref. 7; nt 272–2945 of GenBank accession no. J05121; T686 was changed to A, resulting in substitution of Cys129 by Ser. This was inserted downstream of the SR
promoter). The 4.0 kb SalI fragment containing the SR promoter, p94:C129S coding region and polyadenylation signal was used as the transgene to produce transgenic mice. The fragment was separated by agarose gel electrophoresis, excised from the gel and purified by CsCl ultracentrifugation.
Production, identification and maintenance of transgenic mice
The transgenic mice were produced by microinjection of the linear DNA fragment prepared as above into fertilized eggs of C57BL/6CrSlc mouse (Japan SLC Inc., Shizuoka, Japan) as described previously (43). Genotypes of newborn mice were checked by PCR analyses of the genomic DNA. This was prepared by extracting an ear lobe biopsy (~2 mm diameter) for 1 h at 55°C, in 100 µl of a solution containing 50 mM KCl, 10 mM Tris–HCl buffer pH 8.3, 1.5 mM MgCl2, 0.1% (w/v) gelatin, 0.45% (w/v) NP-40, 0.45% (w/v) Tween-20 and 0.5 mg/ml proteinase K. PCR was performed using the following primers: pSRD-16S, 5'-GCC TGT ACG GAA GTG TTA CTT C-3'; p94-1'A, 5'-CGG AGG TCT CTT CCA GAC GAA CTG-3'; p94-5S, 5'-GAG GAG CAG CAG CAA TTC CGG-3'; p94-5A, 5'-GCA TAC ATG GTA AGC TGC AG-3'.
Presence of the transgene was double checked using two sets of primer pairs, pSRD-16S and p94-1'A and p94-5S and p94-5A. Two microliter samples of the extracted DNA template were amplified in a total of 20 µl of amplification solution, with final concentrations of 0.4 µM each primer, 0.2 mM dNTPs and 0.025 U/µl ExTaq DNA polymerase (Takara Shuzo Co., Kyoto, Japan). The cycling conditions were: 94°C for 30 s; 45 cycles of 94°C for 15 s, 58°C for 60 s, 72°C for 30 s; 72°C for 7 min. The set of primers pSRD-16S and p94-1'A was specific to the transgene, generating a 380 bp PCR fragment, and did not recognize the endogenous p94 gene. Using p94-5S and p94-5A as primers, the transgene was amplified as a 518 bp fragment. The length of the mouse genomic DNA between the positions corresponding to primers p94-5S and p94-5A is ~3 kb. Thus, this region was not amplified with this set of primers under the conditions used. Therefore, these sets of primers distinguished transgenic mice from wild-type individuals. Since the two sets of primers correspond to 5'- and 3'-portions of the transgene, positive bands with both primer sets indicate that nearly the full length was integrated. The founder mice were crossed with wild-type C57BL/6 mice to produce transgenic progeny. Mice were fed on a 12/12 h light/dark cycle at constant temperature (24°C) and humidity (50%), with free access to food and water.
Extraction and analyses of p94 protein from skeletal muscle
Ten mouse skeletal muscle slices (10 µm thickness and 3–4 mm diameter) were homogenized in 100 µl of a solution containing 50 mM KCl, 1 mM EGTA, 5 mM EDTA and 1 mM NaHCO3 and centrifuged at 4°C at 10 000 g for 10 min. The precipitate, which mainly contains connectin (‘connectin fraction’), was then dissolved directly in 20 µl of 1x SDS sample buffer or in 10 µl of phosphate-buffered saline supplemented with 5 µM E-64 and 5 mM EGTA. The latter sample was further incubated on ice or at 37°C for 15 min and the incubation stopped by addition of an equal volume of 2x SDS sample buffer. These samples were separated by SDS–PAGE, transferred onto polyvinylidene difluoride membrane and visualized with p94-specific anti-IS2 rabbit antiserum (27) or affinity-purified anti-IS2 goat antiserum, goat anti-rabbit or horse anti-goat IgG secondary antibody, respectively, and ABC peroxidase kit (Vector Laboratories Inc., Burlingame, CA). The membrane was stained using a peroxidase substrate kit and a POD immunostaining kit (Wako Inc., Osaka, Japan).
Quantification of mRNA by competitive PCR
Total RNA was extracted from skeletal muscle or whole blood cells using Sepasol-RNA II (Nacalai Tesque Inc., Japan) and reverse transcribed using a First-Strand Synthesis kit with a NotI-d(T)18 primer (Amersham Pharmacia Biotech Inc., Piscataway, NJ). DNA competitor was produced using a competitive DNA construction kit, following the manufacturer’s instructions (Takara Shuzo Co.). Briefly, p94:C129S competitor DNA (246 bp) was constructed by PCR to contain the sequence of pSRD-16S at the 5'-end, that of p94-1'A at the 3'-end and 200 bp of 
DNA in the middle. With various amounts of the p94:C129S-competitor, the target template was amplified by PCR using pSRD-16S and p94-1'A as primers under the conditions described above, but for 40 instead of 45 cycles. The detected amount of competitor showed a linear relation with the ratio of the target to the competitor, in a broad concentration range from 1 x 103 to 1 x 106 copies. The initial amount of target was estimated from the point where the ratio of the target to the competitor was one. ß-Actin was also used as a control molecule by making ß-actin competitor DNA (342 bp) similarly to above and performing PCR under the same conditions as above, but for 28 instead of 40 cycles. The primers were 5'-CAG GAG ATG GCC ACT GCC GCA-3' and 5'-TCC TTC TGC ATC CTG TCA GCA-3'. These produce a 275 bp PCR fragment when ß-actin cDNA is amplified.
Evaluation of grip strength
Forelimb grip strength was determined using a Grip Strength Meter MK-380 (Muromachi Kikai Inc., Tokyo, Japan). The mice were allowed to grasp a piece of wire gauze by the forelimbs, and then steadily pulled by the tail horizontally away until the grip was detached. The maximal force values were recorded (44,45). Five such trials were undertaken for each mouse within 2 min and these trials were repeated five times on five different days. Since some lines of a certain age did not contain male mice, we used data only for female mice. The grip strength was normalized as the ratio to body weight and mice of the same age were compared with each other. Statistical analyses were by Student’s t-test.
Histochemical staining of skeletal muscle
Skeletal muscle tissues were frozen in isopentane chilled with liquid nitrogen. Six micrometer serial sections were stained with H&E, mGT or NADH-TR.
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ACKNOWLEDGEMENTS |
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We would like to thank Drs Tatsuya Maeda and Hiroyuki Kawahara for valuable discussions, Dr Takashi Momoi for the generous gift of anti-caspase antibodies, Dr Kenji Wada for introducing us to the grip strength measuring method and Ms Rie Inaba for taking care of the mice. This work was supported in part by a Grant-in-Aid for Scientific Research on Priority Areas, Intracellular Proteolysis, from the Ministry of Education, Science, Sports and Culture, Japan, a Research Grant (11B-1) for Nervous and Mental Disorders from the Ministry of Health and Welfare, Japan, from CREST of the JST and grants from the Kato Memorial Biosciences Foundation and the Human Frontier Science Program.
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FOOTNOTES |
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+ To whom correspondence should be addressed at: Laboratory of Biological Function, Department of Applied Biological Chemistry, Graduate School of Agricultural and Life Sciences, University of Tokyo, 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-8657, Japan. Tel: +81 3 5841 8218; Fax: +81 3 3813 0654; Email: ahsori@mail.ecc.u-tokyo.ac.jp
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REFERENCES |
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