Abstract

Ullrich congenital muscular dystrophy (UCMD) and Bethlem myopathy are inherited muscle disorders caused by mutations of genes encoding the extracellular matrix protein collagen VI (ColVI). Mice lacking ColVI ( Col6a1−/− ) display a myopathic phenotype associated with ultrastructural alterations of mitochondria and sarcoplasmic reticulum, mitochondrial dysfunction with abnormal opening of the permeability transition pore (PTP) and increased apoptosis of muscle fibers. Treatment with cyclosporin (Cs) A, a drug that desensitizes the PTP by binding to cyclophilin (Cyp)-D, was shown to rescue myofiber alterations in Col6a1−/− mice and in UCMD patients, suggesting a correlation between PTP opening and pathogenesis of ColVI muscular dystrophies. Here, we show that inactivation of the gene encoding for Cyp-D rescues the disease phenotype of ColVI deficiency. In the absence of Cyp-D, Col6a1−/− mice show negligible myofiber degeneration, rescue from mitochondrial dysfunction and ultrastructural defects, and normalized incidence of apoptosis. These findings (i) demonstrate that lack of Cyp-D is equivalent to its inhibition with CsA at curing the mouse dystrophic phenotype; (ii) establish a cause–effect relationship between Cyp-D-dependent PTP regulation and pathogenesis of the ColVI muscular dystrophy and (iii) validate Cyp-D and the PTP as pharmacological targets for the therapy of human ColVI myopathies.

INTRODUCTION

Collagen VI (ColVI) is a broadly distributed extracellular matrix protein forming a microfilamentous network that is particularly abundant in skeletal muscle ( 1 , 2 ). The protein is composed of three distinct α-chains encoded by separate genes ( COL6A1 , COL6A2 and COL6A3 in humans). ColVI is synthesized and secreted by cells organizing an extracellular matrix, and transcriptional regulation is a key step in its production. In muscle, ColVI is a major component of the endomysium, where it is localized just outside the basement membrane of muscle fibers ( 3 , 4 ). Deficiency of ColVI in humans, due to mutations in COL6A1-A3 genes, is one of the most common causes of congenital muscular dystrophies ( 1 ). Two main syndromes were described, Bethlem myopathy (MIM#158810) and Ullrich congenital muscular dystrophy (UCMD) (MIM#254090), but additional muscle phenotypes were recently linked to ColVI defects ( 5 ). Bethlem myopathy is a relatively mild and slowly progressive myopathic disorder, whereas UCMD is a severe and rapidly progressive muscle disease usually causing early death due to respiratory failure.

Studies with in vitro cell cultures derived from skeletal muscles have demonstrated that ColVI is largely produced by the interstitial fibroblasts and not by the myogenic cell population ( 4 ). ColVI secreted by muscle interstitial fibroblasts was found to adhere to the surface of myotubes and establish a dense pericellular matrix around myogenic cells ( 4 ). Interestingly, a recent study has shown that transcription of the Col6a1 gene by muscle interstitial fibroblasts is strictly dependent on a specific enhancer region, whose activation requires inductive signals from myogenic cells ( 6 ). These findings suggest that synergistic activities between these two cell types are crucial in regulating ColVI synthesis and function in muscle. Three novel genes ( COL6A4–A6 ) coding for additional ColVI subunits and showing tissue-regulated expression were recently identified ( 7 ), thus increasing the spectrum of potential primary structures for ColVI.

A substantial contribution to understanding the pathogenesis of ColVI diseases was provided by mice lacking ColVI obtained by means of targeted inactivation of Col6a1 gene. We have shown that Col6a1−/− mice have an early onset myopathic phenotype affecting diaphragm and other skeletal muscles, and display spontaneous apoptosis and ultrastructural defects of mitochondria and sarcoplasmic reticulum ( 8 , 9 ). Muscle fibers derived from Col6a1−/− mice have a latent mitochondrial dysfunction due to abnormal opening of the permeability transition pore (PTP) ( 9 ), a high-conductance channel located in the mitochondrial inner membrane that plays a role in several forms of cell death ( 10 ). Consistently, treatment of Col6a1−/− mice with cyclosporin A (CsA), a widely used immunosuppressant that desensitizes the mitochondrial PTP independent of calcineurin inhibition ( 11 ), led to a marked decrease of muscle fiber apoptosis and of ultrastructural lesions ( 9 ). Guided by these findings, we showed that patients affected by UCMD have strikingly similar abnormalities, which could be normalized by treatment with CsA or its non-immunosuppressive derivative Debio 025 ( 12 , 13 ). These observations indicate that the muscle lesions caused by ColVI deficiency can be largely reverted by desensitizing the mitochondrial PTP, yet the mechanistic proof that a cause–effect relationship between PTP opening and the myopathy exits is still lacking.

The mitochondrial target for CsA is cyclophilin (CyP)-D, a peptidyl-prolyl cis–trans isomerase located in the mitochondrial matrix that is inhibited by CsA in the same range of concentrations desensitizing the PTP ( 11 , 14 ). Oxidative and other cellular stresses promote the recruitment of CyP-D to the mitochondrial inner membrane where it favours opening of the PTP ( 10 , 15 ). Mice with targeted inactivation of Ppif , the gene coding for CyP-D, were produced and characterized by different laboratories ( 16–19 ). Mitochondria isolated from Ppif−/− mice show a marked resistance to PTP opening, with a significant desensitization of the PTP to Ca 2+ overload and lack of further desensitizing effects by CsA ( 17 ).

Here, we show that genetic ablation of CyP-D markedly attenuates the phenotypic defects of ColVI-deficient mice. These findings are an important proof of principle that abnormal opening of mitochondrial PTP plays a key mechanistic role in the pathogenesis of ColVI myopathies.

RESULTS

Lack of CyP-D ameliorates muscle histology of Col6a1 knockout mice

Breeding of Col6a1−/− and Ppif−/− mice allowed the generation of animals with either wild-type (WT) ( Col6a1+/+Ppif+/+ ), ColVI null ( Col6a1−/−Ppif+/+ ), CyP-D null ( Col6a1+/+Ppif−/− ) or ColVI/CyP-D double null ( Col6a1−/−Ppif−/− ) genotypes. As reported previously for Ppif−/− mice ( 16–19 ), Col6a1+/+Ppif−/− individuals did not display an overt phenotype, and we verified the lack of muscle abnormalities through histological analysis, electron microscopy and detection of apoptosis (results not shown). Col6a1−/−Ppif−/− mice were born at the expected Mendelian ratios, they were fertile and did not show any overt phenotypic abnormality.

We first performed a histological examination of skeletal muscles from WT, ColVI null and ColVI/CyP-D double null mice. In agreement with previous observations, Col6a1−/− muscles showed typical signs of myopathy, with focal areas of infiltration by phagocytic cells and a pronounced variation of fiber diameter (Fig.  1 Ab and e). These pathological signs were markedly reduced in Col6a1−/−Ppif−/− muscles, which showed more uniform fiber size and decreased tissue inflammation (Fig.  1 Ac and f). Morphometric analysis of myofiber cross-sectional areas in tibialis anterior muscle confirmed that Col6a1−/− contained myofibers of different sizes, with a significant incidence of small fibers, whereas both WT and Col6a1−/−Ppif−/− muscles contained almost exclusively large fibers (Fig.  1 B). Although the amount of myofibers with central nuclei was not very high in ColVI null mice ( 8 , 9 ), Col6a1−/− muscles showed about 8-fold higher incidence of fibers with centrally located nuclei (WT: 0.38 ± 0.16; Col6a1−/− : 3.01 ± 0.71), and this percentage was decreased by half in Col6a1−/−Ppif−/− muscles (1.56 ± 0.03).

Figure 1.

Histological analysis of skeletal muscles from WT, KO ( Col6a1−/− ) and DKO ( Col6a1−/−Ppif−/− ) mice. ( A ) Representative cross (a–c) and longitudinal sections (d–f) of H&E staining of tibialis anterior muscle from 24-week-old WT (a and d), Col6a1−/− (b and e) and Col6a1−/−Ppif−/− (c and f) mice. Col6a1−/− muscles show a large variability in myofiber size, with several fibers of very small diameter (asterisk), presence of regenerating fibers with centrally located nuclei (arrowhead) and occurrence of inflammatory infiltrates (arrows). On the contrary, Col6a1−/−Ppif−/− muscles display uniform myofiber size and absence of inflammatory infiltrates, similarly to WT muscles. Scale bar, 100 µm. ( B ) Morphometric analysis of myofiber cross-sectional areas in tibialis anterior muscles of the different genotypes. Fibers were grouped into three size ranges, and at least 500 fibers of each genotype were analysed using LEICA IM 1000 software (Leica Microsystems). Data are expressed as mean ± SEM. * P < 0.05 versus WT; § not significant ( P = 0.64) versus WT. ( C ) EBD uptake in the diaphragm of WT, Col6a1−/− and Col6a1−/−Ppif−/− mice. Following injection with EBD, diaphragms were isolated from WT (a and a’), KO (b and b’) and DKO (c and c’) mice, and examined by light microscopy (a–c). (a'–c’) Representative cross-sections visualized by fluorescence microscopy, where myofibers penetrated by EBD appear red (arrowheads). Scale bars, 500 µm (a–c) and 100 µm (a'–c’). ( D ) Quantification of EBD-positive fibers in diaphragms of different genotypes. At least five animals of each genotype were analysed, and quantification was carried out in 20 randomly chosen cross-sections covering different areas of the diaphragm. Data are expressed as mean ± SEM. * P < 0.05 versus Col6a1−/− .

Figure 1.

Histological analysis of skeletal muscles from WT, KO ( Col6a1−/− ) and DKO ( Col6a1−/−Ppif−/− ) mice. ( A ) Representative cross (a–c) and longitudinal sections (d–f) of H&E staining of tibialis anterior muscle from 24-week-old WT (a and d), Col6a1−/− (b and e) and Col6a1−/−Ppif−/− (c and f) mice. Col6a1−/− muscles show a large variability in myofiber size, with several fibers of very small diameter (asterisk), presence of regenerating fibers with centrally located nuclei (arrowhead) and occurrence of inflammatory infiltrates (arrows). On the contrary, Col6a1−/−Ppif−/− muscles display uniform myofiber size and absence of inflammatory infiltrates, similarly to WT muscles. Scale bar, 100 µm. ( B ) Morphometric analysis of myofiber cross-sectional areas in tibialis anterior muscles of the different genotypes. Fibers were grouped into three size ranges, and at least 500 fibers of each genotype were analysed using LEICA IM 1000 software (Leica Microsystems). Data are expressed as mean ± SEM. * P < 0.05 versus WT; § not significant ( P = 0.64) versus WT. ( C ) EBD uptake in the diaphragm of WT, Col6a1−/− and Col6a1−/−Ppif−/− mice. Following injection with EBD, diaphragms were isolated from WT (a and a’), KO (b and b’) and DKO (c and c’) mice, and examined by light microscopy (a–c). (a'–c’) Representative cross-sections visualized by fluorescence microscopy, where myofibers penetrated by EBD appear red (arrowheads). Scale bars, 500 µm (a–c) and 100 µm (a'–c’). ( D ) Quantification of EBD-positive fibers in diaphragms of different genotypes. At least five animals of each genotype were analysed, and quantification was carried out in 20 randomly chosen cross-sections covering different areas of the diaphragm. Data are expressed as mean ± SEM. * P < 0.05 versus Col6a1−/− .

We investigated in further detail the histological structure of diaphragm after systemic injection with Evans blue dye (EBD), a small molecular tracer that only permeates the sarcolemma of damaged fibers. Both visual inspection and microscopic analysis showed that WT animals excluded the dye (Fig.  1 Ca and a’), that marked uptake of EBD took place in several fibers in the diaphragm of Col6a1−/− mice (Fig.  1 Cb and b’) and that the number of fibers taking up the dye was remarkably low in the diaphragm of Col6a1−/−Ppif−/− mice (Fig.  1 Cc and c’). Detailed microscopic quantification of EBD-positive fibers in diaphragm transverse sections revealed that 14.6 ± 0.7% of Col6a1−/− muscle fibers were permeable to the tracer, a figure that was reduced to 1.2 ± 0.2% in Col6a1−/−Ppif−/− mice, with a 12-fold decrease compared with Col6a1−/− animals (Fig.  1 D). These data suggest that ablation of CyP-D largely prevents the myopathic defects of ColVI-deficient mice.

CyP-D ablation attenuates ultrastructural defects of ColVI-deficient muscle fibers

Electron microscopic analysis confirmed the characteristic ultrastructural alterations of Col6a1−/− ( 9 ) that are never observed in diaphragms from WT individuals (Fig.  2 A). Col6a1−/− muscle fibers displayed mitochondrial defects with swelling, hypodense matrix and abnormal cristae, and dilations of sarcoplasmic reticulum (Fig.  2 Ab and b’). In Col6a1−/−Ppif−/− mice, lack of CyP-D markedly decreased ultrastructural alterations in diaphragm muscle fibers (Fig.  2 Ac and c’). Analysis of about 300 fibers from three different animals of each genotype showed that Col6a1−/−Ppif−/− mice had a 3-fold lower incidence of fibers with abnormal mitochondria than Col6a1−/− individuals and that this incidence was in the same range of that observed in WT animals (Fig.  2 B). Mitochondria of Col6a1−/−Ppif−/− myofibers typically displayed tightly packed cristae, with a structural organization indistinguishable from that of WT fibers (Fig.  2 A, compare panels a' and c’). Dilations of sarcoplasmic reticulum were still found in Col6a1−/−Ppif−/− muscles (Fig.  2 Ac), but the incidence of fibers with these alterations was lower than that observed in Col6a1−/− muscles, decreasing from 28.1 ± 4.8 to 10.5 ± 5.0% ( P < 0.005).

Figure 2.

( A ) Electron micrographs of diaphragm of WT (a and a’), Col6a1−/− (b and b’) and Col6a1−/−Ppif−/− (c and c’) mice. The pictures show representative fields of whole diaphragm sections (a–c) and mitochondria (a'–c’). In (a–c), enlarged sarcoplasmic reticulum and swollen mitochondria are labelled by arrowheads and asterisks, respectively. Scale bars, 400 nm. ( B ) Frequency of fibers with swollen mitochondria. Data represent the mean of at least three independent experiments ± SD. #P < 0.05 versus WT; * P < 0.05 versus Col6a1−/− ; § not significant ( P = 0.12) versus WT.

Figure 2.

( A ) Electron micrographs of diaphragm of WT (a and a’), Col6a1−/− (b and b’) and Col6a1−/−Ppif−/− (c and c’) mice. The pictures show representative fields of whole diaphragm sections (a–c) and mitochondria (a'–c’). In (a–c), enlarged sarcoplasmic reticulum and swollen mitochondria are labelled by arrowheads and asterisks, respectively. Scale bars, 400 nm. ( B ) Frequency of fibers with swollen mitochondria. Data represent the mean of at least three independent experiments ± SD. #P < 0.05 versus WT; * P < 0.05 versus Col6a1−/− ; § not significant ( P = 0.12) versus WT.

Lack of CyP-D prevents mitochondrial depolarization in myofibers and cultured muscle cells of ColVI-deficient mice

We next assessed the mitochondrial membrane potential by monitoring the accumulation of tetramethylrhodamine methyl ester (TMRM) in flexor digitorum brevis (FDB) muscle fibers derived from mice with the different genotypes. We previously demonstrated that Col6a1−/− myofibers display an anomalous depolarizing response elicited by the addition of oligomycin, an inhibitor of the mitochondrial F1FO ATPase ( 9 ). As expected, the addition of oligomycin caused higher incidence of fibers with depolarizing mitochondria in Col6a1−/− compared with WT mice, the fraction of fibers with depolarizing mitochondria after oligomycin addition being, respectively, 33 and 10% for these two genotypes (Fig.  3 A, left and middle panels). In Col6a1−/−Ppif−/− individuals, lack of CyP-D led to an increased resistance to oligomycin-induced mitochondrial depolarization, and the fraction of fibers with depolarizing mitochondria dropped to 14% (Fig.  3 A, right panel). Our previous studies on isolated myofibers and primary muscle cell cultures indicated that the deficiency of extracellular ColVI results in an increased open propensity of the PTP, thus impinging on the mitochondrial ability to maintain the membrane potential in response to various treatments ( 9 , 12 , 20 ). Based on these observations, we monitored TMRM fluorescence in primary diaphragm cell cultures derived from mice with the three genotypes, in order to assess how ablation of CyP-D affected mitochondrial membrane potential in different experimental conditions. A marked decrease of TMRM fluorescence was induced by either oligomycin or rotenone in primary muscle cultures from Col6a1−/− mice, but not in those from WT and Col6a1−/−Ppif−/− animals (Fig.  3 B and C). Addition of the protonophore carbonylcyanide- p -trifluoromethoxyphenyl hydrazone promptly collapsed fluorescence in all cultures, thus confirming that the observed variations in TMRM fluorescence were indeed measuring the mitochondrial membrane potential. The responses elicited by oligomycin and rotenone in WT and Col6a1−/−Ppif−/− cells were remarkably similar, and both cultures were able to maintain high TMRM fluorescence after more than 1 h of incubation in the presence of either inhibitor (Fig.  3 B and C).

Figure 3.

( A ) Mitochondrial response to oligomycin in muscle fibers of WT ( n = 30), Col6a1−/− (KO, n = 54) and Col6a1−/−Ppif−/− (DKO, n = 30) mice. Myofibers were isolated from FDB muscle and loaded with TMRM (20 n m ) for 15 min at 37°C. Where indicated (arrows), oligomycin (Oligo, 5 µ m ) or the protonophore carbonyl cyanide p- trifluoromethoxyphenylhydrazone (FCCP, 4 µ m ) were added. The probe accumulates in mitochondria that maintain the membrane potential. Results of at least three experiments per genotype are shown, and each trace represents the fluorescence values of a single fiber. The fraction of depolarizing fibers (as per cent of total) is also provided for each genotype, where fibers are considered as depolarizing when they lose more than 10% of initial value of TMRM fluorescence after oligomycin addition. ( B and C ) Changes of mitochondrial TMRM fluorescence induced by oligomycin (B) or rotenone (C) in primary muscle cell cultures derived from diaphragms. Cells from WT (open circles), Col6a1−/− (closed squares) and Col6a1−/−Ppif−/− (open squares) were loaded with 10 n m TMRM for 30 min at 37°C. Where indicated, oligomycin (Oligo, 5 µ m ), rotenone (Rot, 2 µ m ) or carbonylcyanide- p- trifluoromethoxyphenylhydrazone (FCCP, 4 µ m ) were added. Data represent the mean of at least four independent experiments ± SEM. The upper bars indicate all the time points where P < 0.05 for Col6a1−/− compared with either WT (*) or with Col6a1−/−Ppif−/− (#).

Figure 3.

( A ) Mitochondrial response to oligomycin in muscle fibers of WT ( n = 30), Col6a1−/− (KO, n = 54) and Col6a1−/−Ppif−/− (DKO, n = 30) mice. Myofibers were isolated from FDB muscle and loaded with TMRM (20 n m ) for 15 min at 37°C. Where indicated (arrows), oligomycin (Oligo, 5 µ m ) or the protonophore carbonyl cyanide p- trifluoromethoxyphenylhydrazone (FCCP, 4 µ m ) were added. The probe accumulates in mitochondria that maintain the membrane potential. Results of at least three experiments per genotype are shown, and each trace represents the fluorescence values of a single fiber. The fraction of depolarizing fibers (as per cent of total) is also provided for each genotype, where fibers are considered as depolarizing when they lose more than 10% of initial value of TMRM fluorescence after oligomycin addition. ( B and C ) Changes of mitochondrial TMRM fluorescence induced by oligomycin (B) or rotenone (C) in primary muscle cell cultures derived from diaphragms. Cells from WT (open circles), Col6a1−/− (closed squares) and Col6a1−/−Ppif−/− (open squares) were loaded with 10 n m TMRM for 30 min at 37°C. Where indicated, oligomycin (Oligo, 5 µ m ), rotenone (Rot, 2 µ m ) or carbonylcyanide- p- trifluoromethoxyphenylhydrazone (FCCP, 4 µ m ) were added. Data represent the mean of at least four independent experiments ± SEM. The upper bars indicate all the time points where P < 0.05 for Col6a1−/− compared with either WT (*) or with Col6a1−/−Ppif−/− (#).

CyP-D inactivation prevents muscle apoptosis in ColVI-deficient mice

We finally investigated whether genetic ablation of CyP-D was able to improve cell survival and to counteract the apoptotic phenotype observed in Col6a1−/− mice ( 9 ). The frequency of apoptotic nuclei was evaluated in situ by using the terminal deoxynucleotidyl transferase-mediated dUTP nick end labelling (TUNEL) method. TUNEL-positive nuclei were readily detected in the diaphragm of Col6a1−/− mice, but not in the corresponding samples of WT and Col6a1−/−Ppif−/− mice (Fig.  4 A). Col6a1−/− muscle sections displayed an average of 57.0 ± 3.8 TUNEL-positive nuclei/mm 2 , while the incidence in Col6a1−/−Ppif−/− sections was 4.8 ± 1.6, which is not significantly different from the value (2.0 ± 1.2) measured in WT sections (Fig.  4 B). To investigate further the link between CyP-D ablation and increased cell survival, we also assessed the occurrence of apoptosis in cultured muscle cells. Again, cultures from Col6a1−/−Ppif−/− and WT muscles displayed similar frequencies of TUNEL-positive nuclei, and these were significantly lower than those displayed by cultures from Col6a1−/− muscles (Fig.  4 C). Growth of Col6a1−/−Ppif−/− cultures in serum-free medium caused a 5-fold increase in the incidence of TUNEL-positive nuclei; as expected of cells lacking CyP-D, however, the apoptotic response to serum starvation was not prevented by treatment with CsA (Fig.  4 C).

Figure 4.

Incidence of apoptosis in muscle ( A and B ) and in cell cultures ( C ) derived from WT, Col6a1−/− and Col6a1−/−Ppif−/− mice. (A) TUNEL staining of diaphragms. Hoechst-stained sections are shown below the corresponding TUNEL pictures. Several TUNEL-positive nuclei (arrowheads) are present in KO, but not in WT and DKO muscles. Scale bar, 50 µm. (B) Incidence of TUNEL-positive nuclei in diaphragm muscle fibers with the indicated genotype. Data are expressed as mean ± SEM of at least three independent experiments. For each genotype, 20–30 animals were analysed. * P < 0.05 versus Col6a1−/− ; § not significant ( P = 0.13) versus WT. (C) Primary muscle cells were derived from diaphragms of mice with the stated genotypes, cultured on plastic dishes and scored for the presence of TUNEL-positive nuclei. Where indicated, cells were maintained for 2 h in serum-free (NS) medium in the absence or presence of 1.6 µ m CsA. Data are the mean of four independent experiments ± SEM. * P < 0.05 versus Col6a1−/− ; § not significant ( P = 0.41) versus WT; §§ not significant ( P = 0.93).

Figure 4.

Incidence of apoptosis in muscle ( A and B ) and in cell cultures ( C ) derived from WT, Col6a1−/− and Col6a1−/−Ppif−/− mice. (A) TUNEL staining of diaphragms. Hoechst-stained sections are shown below the corresponding TUNEL pictures. Several TUNEL-positive nuclei (arrowheads) are present in KO, but not in WT and DKO muscles. Scale bar, 50 µm. (B) Incidence of TUNEL-positive nuclei in diaphragm muscle fibers with the indicated genotype. Data are expressed as mean ± SEM of at least three independent experiments. For each genotype, 20–30 animals were analysed. * P < 0.05 versus Col6a1−/− ; § not significant ( P = 0.13) versus WT. (C) Primary muscle cells were derived from diaphragms of mice with the stated genotypes, cultured on plastic dishes and scored for the presence of TUNEL-positive nuclei. Where indicated, cells were maintained for 2 h in serum-free (NS) medium in the absence or presence of 1.6 µ m CsA. Data are the mean of four independent experiments ± SEM. * P < 0.05 versus Col6a1−/− ; § not significant ( P = 0.41) versus WT; §§ not significant ( P = 0.93).

DISCUSSION

The results of the present manuscript provide compelling evidence for the mitochondrial pathogenesis of ColVI myopathies, which we had previously suggested based on ultrastructural, functional and pharmacological data ( 9 , 12 , 13 , 20 ). Mitochondrial dysfunction can be readily demonstrated in skeletal muscle fibers and muscle-derived cultures both from Col6a1−/− mice and from patients affected by UCMD ( 9 , 12 , 13 , 20 ), as also recently confirmed by another laboratory ( 21 ). These findings, however, as well as the striking normalizing effect of CsA on mitochondrial function ( 9 , 12 , 13 , 20 , 21 ) and apoptosis ( 9 , 12 , 13 , 20 ) provided an intriguing correlation but not the proof that a cause–effect relationship exists. Indeed, two major issues should be considered when interpreting the pharmacological effects of CsA on ColVI myopathies.

CsA also inhibits calcineurin, which has been shown to affect skeletal muscle physiology ( 22 ) and mitochondrial fission through Drp-1 dephosphorylation ( 23 ). We had previously addressed this issue in the Col6a1−/− mouse and found that inhibition of calcineurin with FK506, which does not inhibit CyPs and has no effects on the PTP ( 24 ), could not reproduce the protective effects of CsA but rather worsened the incidence of apoptosis ( 9 ). However, it is difficult to extrapolate the effects obtained in the mouse model to human ColVI diseases, which are genetically and functionally so heterogeneous ( 1 ). A strong indication that calcineurin is not involved in the effects of CsA in the patients was obtained through the use of D-MeAla 3 -EtVal 4 -cyclosporin (Debio 025), a CyP inhibitor that does not affect calcineurin. Indeed, Debio 025 normalized the mitochondrial phenotype and decreased the incidence of apoptosis in cultures from UCMD patients ( 12 ) and Col6a1−/− mice ( 44 ); yet it was 7000 times less active than CsA at inhibiting interleukin-2 production in Jurkat cells and at least 15 times less potent in mixed lymphocyte reaction tests ( 25 ).

Even if a role for calcineurin inhibition in the protective effects of CsA could be ruled out in ColVI deficiencies, a second problem is that CsA (and Debio 025) binds to and inhibits all members of the CyP family of peptidyl-prolyl cis–trans isomerases ( 26 ), which in humans includes 17 unique proteins ( 27 ). Besides their possible implication in ColVI myopathies, CyPs play important roles in a number of human diseases that include inflammation and vascular dysfunction ( 28–32 ), wound healing ( 33 ), innate immunity to HIV ( 34 ), hepatitis C infection ( 35 ), host–parasite interactions ( 36 ), tumour biology ( 37 ) and several pathological conditions mediated by the mitochondrial PTP through matrix CyP-D ( 10 ). Thus, and despite the demonstrable beneficial effects of CsA and Debio 025 on mitochondria in ColVI diseases ( 9 , 12 , 13 , 20 ), it is impossible to sort the consequences of CyP-D inhibition from those caused by inhibition of other CyPs, which might independently affect muscle fiber apoptosis. Thus, assessing whether a cause–effect relationship exists between PTP inhibition and therapy of ColVI muscular dystrophies remained a major challenge that could only be addressed by genetics.

In the present study, we took advantage of a CyP-D null mouse strain that we had developed and characterized previously ( 17 , 38 ). Mitochondria isolated from these mice display a decreased sensitivity of the PTP to opening that matches quite precisely the effects of CsA in WT individuals ( 17 ) provided that inorganic phosphate is present ( 38 ). Despite lack of an overt phenotype, CyP-D null mice proved to be extremely resistant to ischaemia-reperfusion injury of the heart ( 16 , 18 ) and brain ( 19 ). Crossing of these mice with the Col6a1−/− mouse ( 8 ) offered, therefore, a unique opportunity to test whether genetic ablation of CyP-D could mimic the protective effects of CsA on the myopathy.

The striking result of the present work is that Col6a1−/−Ppif−/− mice no longer display the disease phenotype of Col6a1−/− individuals, which is characterized by histological features of myopathy, permeability of skeletal muscle fibers to small tracers ( 8 ), ultrastructural lesions of the mitochondria and sarcoplasmic reticulum ( 9 ), latent mitochondrial dysfunction that can be triggered by oligomycin ( 9 ) or rotenone ( 20 ) and increased rates of apoptosis ( 9 ). With the exception of a small, residual dilation of the sarcoplasmic reticulum, all these hallmarks of the ColVI disease disappeared in Col6a1−/−Ppif−/− mice. This set of findings demonstrates that genetic ablation of CyP-D, a molecule that favours PTP opening by shielding it from the protective effects of inorganic phosphate ( 38 ), has the same beneficial effects on the myopathic phenotype caused by ColVI deficiency as pharmacological inhibition of CyP-D with CsA or Debio 025 ( 9 , 12 , 13 , 20 ). Our findings match recent genetic and pharmacological studies that extend the role of the PTP to mouse models of Duchenne muscular dystrophy ( 39 , 40 ) and MDC1A ( 39 ).

Taken together, these results have tremendous implications for the therapeutic perspectives of muscular dystrophies, in general, and of ColVI diseases, in particular. UCMD is a chronic muscle wasting disease involving the diaphragm, and respiratory failure is a common complication which is worsened by pulmonary infections ( 41 ). Long-term treatment with CsA exposes the patients to the untoward effects of immunosuppression, which may favour life-threatening infections. The present identification of the PTP as a causative event in muscle cell demise in ColVI muscular dystrophy and the genetic validation of CyP-D as a calcineurin-independent pharmacological target represent a fundamental step towards a therapy of human ColVI muscular dystrophies with CyP inhibitors.

MATERIAL AND METHODS

Mice

Col6a1−/− mice were crossed with Ppif−/− mice. Both strains were previously characterized and were in the inbred C57BL/6 background ( 8 , 17 ). All experiments were performed by comparing sex matched, 8- to 24-week-old animals with either Col6a1−/−Ppif+/+ [ColVI knockout (KO)], Col6a1−/−Ppif−/− [ColVI/CyP-D double knockout (DKO)] or Col6a1+/+Ppif+/+ (WT) genotypes. Mouse procedures were approved by the competent Authority of the University of Padova and authorized by the Italian Ministry of Health.

Histology and EBD uptake

Tibialis anterior muscles were removed, frozen in isopentane, cooled in liquid nitrogen and kept at −80°C until use. Longitudinal and cross-sections (7 μm) were prepared and processed for haematoxylin and eosin (H&E) staining. EBD staining of muscles in vivo was performed by i.p. injection with 0.2 ml EBD (10 mg/ml in phosphate-buffered saline, Sigma). Mice were sacrificed after 16–18 h and diaphragms were fixed with 4% paraformaldehyde overnight at 4°C, dehydrated in a graded series of water/ethanol mixtures and embedded in paraffin. Seven-micrometre-thick sections were cut and examined with a Zeiss Axioplan fluorescence microscope (20× magnification) to locate EBD distribution after the removal of paraffin and mounting in Permount.

Electron microscopy

Diaphragm muscles were isolated from mice, gently stretched on wax, fixed with 2.5% glutaraldehyde in phosphate buffer 0.1 m (pH 7.4), postfixed with 1% osmium tetroxide in veronal buffer and embedded in Epon E812 resin. Ultrathin sections were stained with uranyl acetate and lead citrate, and observed in a Philips EM400 transmission electron microscope at 100 kV. For statistical analysis, 300 myofibers were observed from three different tissue blocks for each sample. Data were analysed with the Mann–Whitney test, and values with P < 0.05 were considered significant.

Isolation of skeletal muscle fibers

Fibers were isolated from FDB muscles as previously described ( 9 , 42 ). Intact myofibers were plated onto 24 mm round glass coverslips coated with laminin (3 µg cm −2 ) and cultured for 24 h in Dulbecco’s modified Eagle medium (DMEM, Sigma) containing 10% fetal calf serum (FCS, Sigma), before starting the experiment.

Primary muscle cell cultures

Cultures enriched in myoblasts were prepared by enzymatic and mechanical dissociation of diaphragms from 8-week-old mice, using a protocol modified from Rando and Blau ( 43 ). Briefly, the dissected muscles were minced and incubated for 40 min at 37°C in DMEM supplemented with 0.5% collagenase type I (Sigma). Cells released from the tissue were passed through a 80 μm filter, collected by centrifugation and resuspended in DMEM supplemented with 20% FCS. The cell suspension was plated in culture dishes and incubated at 37°C for 15 min, to allow for adhesion of fibroblasts. Floating cells, consisting of an enriched myoblast population, were transferred to gelatin-coated dishes and grown at 37°C and 5% CO 2 in DMEM supplemented with 20% FCS. Cultures were characterized as described in the supplemental material ( Supplementary Material, Figs S1 and S2; and Table S1 ), and the relative proportion of myoblasts versus fibroblasts was carefully determined. Primary cultures within passage 6 were used for all experiments.

TMRM assay

Mitochondrial membrane potential was measured by epifluorescence microscopy based on the accumulation of TMRM fluorescence. FDB myofibers were placed in 1 ml Tyrode’s buffer and loaded with 20 n m TMRM (Molecular Probes) as previously described ( 9 ). Primary muscle cells cultures were seeded onto 24 mm-diameter round glass coverslips, grown for 2 days in DMEM supplemented with 20% FCS and studied as described after loading with 10 n m TMRM in 1 ml of serum-free DMEM ( 12 ).

TUNEL assay

Paraffin sections (7 µm thick) were prepared from diaphragm muscles, after fixation with 4% paraformaldehyde and paraffin embedding. TUNEL was performed with the ApopTag peroxidase in situ apoptosis detection system (Chemicon). Visualization of all nuclei was obtained by staining with Hoechst 33258 (Sigma). The number of total and TUNEL-positive nuclei for each genotype was determined in at least 300 randomly selected fields by using a Zeiss Axioplan microscope (40× magnification) equipped with a digital camera.

Statistical analysis

Except where indicated, data were analysed with the unpaired Student’s t- test, and values with P < 0.05 were considered significant. Data are presented as mean ± SEM or mean ± SD.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at HMG online.

FUNDING

This study was supported by Telethon-Italy (Grant GGP08107), the Italian Ministry for University and Research and the Association Française contre les Myopathies.

ACKNOWLEDGEMENT

We thank Anna Urciuolo for help with the collagen VI immunostaining.

Conflict of Interest statement . None declared.

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in press

Author notes

These authors contributed equally to this work.