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(American Journal of Pathology. 2003;162:1001-1009.)
© 2003 American Society for Investigative Pathology

Changes in Myotonic Dystrophy Protein Kinase Levels and Muscle Development in Congenital Myotonic Dystrophy

Denis Furling^*, Le Thanh Lam, Onnik Agbulut^*, Gillian S. Butler-Browne^* and Glenn E. Morris

From the Centre National de la Recherche Scientifique Unité Mixte de Recherche (CNRS UMR) 7000,^* Faculté de Médecine Pitié-Salpêtrière, Paris, France; and the Biochemistry Group, North East Wales Institute, Wrexham, United Kingdom

	Abstract

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Myotonic dystrophy (DM1) is caused by the expansion of a CTG repeat in the noncoding region of a protein kinase, DMPK, expressed in skeletal and cardiac muscles. The aim of the present study was to determine the effects of very large CTG expansions on DMPK expression and skeletal muscle development. In fetuses suffering from the severe congenital form of DM1 with large CTG expansions (1800 to 3700 repeats), the skeletal muscle level of DMPK was reduced to 57% of control levels and a similar reduction was observed in cultured DM1 muscle cells relative to control cultures. These results are consistent with greatly reduced DMPK expression from the mutant allele and normal expression from the unaffected allele in this autosomal dominant disorder. In normal fetuses, DMPK protein levels increased dramatically between 9 and 16 weeks and remained high throughout the remaining gestation period. DM1 fetuses showed impaired skeletal muscle development, characterized by a persistence of embryonic and fetal myosin heavy chains and almost total absence of slow myosin heavy chains at the end of gestation. DMPK expression, however, was similar in both fast and slow fibers from normal adult muscle. The reduced DMPK and the delayed slow fiber maturation in congenital DM1 may be two separate consequences of nuclear retention of DMPK RNA transcripts with expanded CUG repeats.

In skeletal muscle, the most common features in the adult or classic form of the disease are myotonia and muscle wasting. The main clinical symptoms of the most severe form of DM1, congenital myotonic dystrophy (CDM) are hypotonia and neonatal respiratory distress, the latter being the major cause of mortality in affected infants. Mental retardation and delayed motor milestones are frequently observed in children that survive. CDM is usually associated with extremely large CTG expansions (>1500 CTG) and evidence of delayed or arrested muscle maturation.^9,10

Various hypotheses have been proposed to explain how this untranslated CTG repeat causes the multitude of physiopathological features of DM. Experimental support has been produced both for dominant-negative effects of CUG repeats in RNA transcripts and for haploinsufficiency due to reduced expression of DMPK and its neighboring genes. Nuclear retention of DMPK transcripts with expanded CUG repeats has been demonstrated¹¹ and decreased cytoplasmic DMPK mRNA levels have been reported in DM tissues.^12,13 This is consistent with either model. Recently, however, DM2 (or PROMM), a disease with very similar clinical features to DM1, was shown to be caused by large CCTG repeat expansions in an unrelated region of the human genome.¹⁴ In both DM1 and DM2, mutant transcripts are retained in the nucleus as discrete foci, implying that dominant-negative effects of expanded repeats in nuclear RNA may be the primary pathogenic mechanism in both diseases. The expanded CUG repeat in DM1 can affect the levels of CUG-binding proteins in the nucleus and change the alternative splicing pattern of several important RNA transcripts.^15,16 Furthermore, human transcription factors related to Drosophila muscleblind bind only to expanded repeats and are sequestered into nuclear foci containing CUG repeats in both DM1 and DM2,^17,18 with possible consequences for muscle development. Although the nuclear RNA hypothesis may seem to minimize the importance of the host genes that contain the repeats, studies with knockout mice suggest that reduced levels of DMPK in heterozygote mice can cause cardiac conduction defects similar to those observed in DM1.¹⁹ Some of the skeletal muscle features of DM are also observed in DMPK knockout mice²⁰ and cataracts are observed in Six5 knockout mice and heterozygotes.^21,22 Unless the knockout phenotypes are fortuitous, these studies suggest that haploinsufficiency of DMPK and Six5 may contribute to the DM1 phenotype, in addition to the dominant-negative effects of the nuclear repeats. In contrast, the characteristic myotonia of DM is only reproduced in mice overexpressing long CUG repeats in muscle nuclear RNA.^23,24

The human DMPK cDNA has 15 exons and predicts a protein of 70 kd.^1,2,25 DMPK is a cyclic AMP-dependent protein kinase with a large catalytic domain followed by smaller coiled-coil and hydrophobic C-terminal domains. Alternative splicing has been observed in a short VSGGG sequence between domains and in the C-terminal domain.²⁶ Recent studies suggest that authentic DMPK migrates with an apparent Mr of 80 to 85 kd on standard sodium dodecyl sulfate-polyacrylamide gel electrophoresis.²⁷ Earlier studies suggesting various Mrs of between 45 kd and 72 kd^28-39 may have been using antibodies that were not entirely specific for DMPK. Some of the proteins that cross-react with the anti-DMPK antibodies^27,40 have been identified as isoforms of MRCK (myotonic dystrophy-related cdc42-binding kinase,^41,42 the protein most closely related to DMPK in sequence. Previous comparisons of DM patients with controls have either measured DMPK-like proteins of the wrong Mr or, in studies that included an 80- to 85-kd DMPK, produced conflicting results.

One aim of the present study was to determine whether nuclear retention of DMPK mRNA in DM1 results in a reduction in DMPK levels, as predicted for complete retention, or whether up-regulation of DMPK protein production from the unaffected allele occurs as a compensation mechanism. We chose to study DMPK levels in congenital DM1 muscles with very large CTG repeats because nuclear retention of DMPK mRNA from the affected allele is likely to be maximal, if not complete. A second aim was to produce a definitive study of fast and slow fiber maturation in DM1 by measuring embryonic, fetal, fast and slow isoforms of myosin heavy-chain and to determine whether preferential slow fiber loss in DM1 is related to preferential DMPK expression in slow fibers, as suggested by studies with polyclonal anti-DMPK sera.^43-45

	Materials and Methods

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Materials

Biopsies from quadriceps muscles were obtained during autopsies, in accordance with French legislation on ethical rules. Fetal muscles were taken for diagnostic purposes from therapeutic abortuses (up to 26 weeks) or from premature or caesarian stillbirths (after 31 weeks). DM fetuses of 16 and 19 weeks were aborted after molecular diagnosis of severe DM (CTG repeats of 2000 and 1800). The 26-week DM fetus was hypotrophic with reduced fetal movements and talipes and died at birth. The 31 week (caesarian) and 33-week DM fetuses died of respiratory distress shortly after birth. The control biopsies (8, 9, 16, 18, 19, 22, 25, 33, and 35 weeks) were from aborted fetuses showing no sign of neuromuscular disease.

Human Skeletal Muscle Cell Culture and Muscle Fiber Isolation

Satellite cells were isolated from normal and CDM fetal muscle samples as described previously.⁴⁶ Myoblasts were grown in Ham’s F10 medium (Gibco, Paisley, UK) supplemented with 50 µg/ml of gentamicin (Biomedia, Boussens, France) and 20% fetal calf serum (Biomedia). All cultures were incubated at 37°C in a humid air atmosphere containing 5% CO₂. For myoblast differentiation, growth medium was removed from subconfluent cultures and replaced by Dulbecco’s modified Eagle’s medium (Gibco) containing 10 µg/ml of insulin and 100 µg/ml of transferrin (Sigma, St Louis, MO). Human muscle fibers were isolated as described by Bonavaud and colleagues.⁴⁷ Briefly, enzymatic digestion of the muscle with collagenase (0.2%) was performed and single fibers were isolated by repeatedly triturating the muscle fragment with a Pasteur pipette. Only intact isolated fibers were selected.

Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis and Western Blotting

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis and Western blotting were performed essentially as described elsewhere,⁴⁸ using 7% or 10% polyacrylamide gels. A panel of previously described monoclonal antibodies against DMPK²⁷ were used at a 1/100 dilution of the culture supernatant for Western blots. Analysis of the myosin heavy chain (MyHC) isoforms was performed using different monoclonal antibodies that react specifically with the embryonic (F1652; Biovalley, Conches, France), fetal, slow and fast (Novocastra, Newcastle, UK) MyHC, or with all of the sarcomeric MyHCs (MF-20⁴⁹ ). Monoclonal antibodies against dystrophin,⁵⁰ emerin,⁵¹ ß-dystroglycan,⁵² and actin (ac-74, Sigma) were also used as controls. Antibody-reacting bands were visualized after development with peroxidase-labeled horse anti-mouse Ig (1/1000; Vector Laboratories, Burlingame, CA) and a chemiluminescent detection system (SuperSignal; Pierce, Rockford, IL).

For quantitation, images of Coomassie Blue-stained gels and developed X-ray films from Western blots were captured using a video camera and frame-grabber. Image analysis was performed using Laserpix software (BioRad Laboratories, Hemel Hempstead, UK), which compensates for nonlinear optical density responses. A measure of the relative protein content of stained gel lanes was determined from the sum of the peak areas of the three major protein bands (myosin, actin, and tropomyosin) that could be measured accurately. DMPK peak areas were divided by this protein peak area value to obtain an estimate of relative DMPK concentration in arbitrary units. Comparative measurements were made within a single gel or Western blot, because it is not possible to compare optical densities between blots. A gel with different sample dilutions was used to confirm a linear response over the range of optical densities observed.

RNA Extraction and Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR)

Total cellular RNA was isolated from skeletal muscle tissues with Trizol reagent (InVitrogen, Cergy, Pontoise, France) according to the manufacturer’s protocol. First-strand cDNA was synthesized using equal amounts of total RNA (5 µg) in a reaction mixture of 200 U of M-MLV reverse-transcriptase (InVitrogen), 1x RT-PCR reaction buffer, 10 mmol/L each dNTPs, 0.1 mol/L of dithiothreitol, 50 U of RNasin (Promega, Madison, WI), 0.1 µg of random hexamers, and 0.1 µg oligo(dT) at 42°C for 1 hour. Subsequently, one-fifth of RT reaction was used as a template for PCR analyses. Standard PCR reaction mixture conditions containing 250 µmol/L dNTPs, 2.5 U of TaqDNA polymerase (Qiagen, Valencia, CA), 1x PCR reaction buffer and 200 ng of upstream and downstream primers (specific for the insulin receptor (InR) and described previously¹⁶ ) were used. Thirty-three cycles of amplification were performed, each consisting of 1 minute at 94°C, 1 minute at 60°C, and 1 minute at 72°C, followed by a final 10-minute extension at 72°C. PCR amplification of insulin receptor mRNA (fragments of 167 bp for the B isoform and 131 bp for the A isoform) were resolved on 3% agarose gel slabs stained with ethidium bromide.

	Results

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DMPK levels were estimated by Western blotting of total skeletal muscle extracts from different stages of human development and from human satellite cell cultures, using a panel of DMPK-specific monoclonal antibodies.²⁷ The principal mAbs used in this study were MANDM13 and MANDM9 (directed against the coil domain) and MANDM4 (directed against the catalytic domain), all of which were specific for the 80- to 85-kd DMPK. MANDM1 (directed against the catalytic site), recognizes the 80- to 85-kd DMPK but cross-reacts with at least two other proteins, including MRCK. Most experiments were done with all mAbs and representative results with some of them are shown.

DMPK Expression Increases during Skeletal Muscle Differentiation in Vitro

As shown recently, DMPK is expressed almost exclusively in muscle and heart.²⁷ We measured the levels of DMPK during the myogenic differentiation of human skeletal muscle cells in culture and results using both catalytic (MANDM1) and coil domain (MANDM9) mAbs are shown. By switching the cultures from permissive to nonpermissive conditions, the muscle cells are able to fuse and form multinucleated myotubes. Figure 1 shows that DMPK was already present at low levels before changing the culture medium and increased approximately fivefold between day 0 and 8 of myogenic differentiation. The levels of a 72-kd cross-reacting protein, CRP, recognized only by MANDM1 mAb, also increased fivefold during the same period of differentiation. MRCK, which is mainly a brain protein, did not increase significantly.

View larger version (43K): [in this window] [in a new window]   Figure 1. The expression of DMPK and related proteins during the differentiation of human skeletal muscle cells in vitro. Muscle cell cultures were grown in differentiation medium for 10 days. DMPK (80 to 85 kd) levels were determined using MANDM9 and MANDM1, whereas MRCK (190 to 200 kd) and CRP (72 kd) levels were determined with MANDM1 alone (insets). Corrections for loading were applied by densitometry of a Coomassie Blue-stained gel using the average of three minor protein bands (inset: correction ratios for 0:1:4:8 days were 1:0.96:2.08:1.42). Major protein bands, such as actin and myosin, were avoided for this correction because myofibrillar proteins increase greatly during myogenesis. For DMPK, the average value from the two mAbs was used to construct the curve. The protein band on the Western blot between MRCK and DMPK has not been identified and the possibility of an MRCK degradation product has not been ruled out. Traces of a protein band below DMPK on the MANDM9 Western blot may also be a degradation product of DMPK.

DMPK levels were determined in fetal, newborn, and adult muscles and results with mAbs against both catalytic (MANDM4) and coil (MANDM13) domains are shown in Figure 2 . DMPK was not detected in fetal muscle extracts at 8 and 9 weeks of development when primary myotubes are just beginning to form but was present at 16 and 18 weeks at the end of secondary myotube formation (Figure 2A) . Control mAbs against emerin or ß-dystroglycan were used to show that sufficient total protein had been loaded for the 8- and 9-week samples. To follow the expression and the levels of DMPK in muscle during human development, MANDM13 mAb was used to quantitate DMPK by microdensitometry of Western blots. Equal amounts of each extract were loaded as far as possible but because this could not be done with complete accuracy, corrections were also applied for protein loading by microdensitometry of Coomassie Blue-stained gels. Several different protein bands, including actin and myosin, were used for normalization, but the final results were similar. Figure 2B shows that DMPK levels remained high from 18 weeks of fetal development until 10 days postnatally. There was an apparent decline to 40% of these levels in adult muscle (32+ years) and the levels of CRP also decreased by 50% (Figure 2B) , but larger numbers of adult samples would be required to determine whether these reductions are statistically significant.

View larger version (43K): [in this window] [in a new window]   Figure 2. A: The expression of DMPK in developing human skeletal muscle. The Western blots were developed using MANDM4 or MANDM13 mAb,26 as indicated, for DMPK at 80 to 85 kd. The same muscle extracts were developed with control mAbs for emerin (MANEM549 ) or ß-dystroglycan (MANDAG250 ) to demonstrate approximately equal loading of protein in each extract. Emerin levels change little during development49 and ß-dystroglycan is present in all cells, although it does increase during myogenesis (see 8- and 16-week lanes). B: DMPK and CRP levels at different stages of human muscle development. Gel lanes were loaded with approximately equal amounts of total protein and Western blots were incubated with MANDM1 mAb, which recognizes both DMPK at 80 to 85 kd and a cross-reacting protein (CRP at 72 kd). The inset shows the Western blot (f, fetal; w, weeks; +, postnatal; y, years). The blot bands were quantified and corrected for loading by quantification of a parallel Coomassie Blue-stained gel as described in Materials and Methods.

To determine whether DMPK is expressed specifically in fast or slow or in both types of fibers of human skeletal muscle, we isolated single muscle fibers from an adult skeletal muscle. Expression of slow or fast MyHC isoforms was used to characterize the phenotype of the single fibers. Figure 3 shows that only slow MyHC (and no fast MyHC) was detected in the slow fiber, whereas the fast fiber expresses only fast MyHC. However, DMPK was detected in both fast and slow fibers. Figure 3B shows immunolocalization of DMPK in fixed adult human sections (Figure 3B, a) . These sections contained both fast and slow muscle fibers in approximately equal abundance (Figure 3B, c) . All fibers stained uniformly in the sarcoplasm by the DMPK mAb, confirming the Western blot evidence in Figure 3A for the presence of DMPK at similar levels in both fiber types.

View larger version (49K): [in this window] [in a new window]   Figure 3. DMPK is expressed in both slow and fast muscle fibers. A: Single muscle fibers were isolated from adult muscle as described in the Materials and Methods section and characterized using the expression of slow or fast MyHC. DMPK was detected in both fast and slow fibers by Western blot using MANDM9. B: Human muscle frozen sections were treated with 3% formalin in PBS for 10 minutes and blocked with 1 mol/L of glycine in PBS. They were incubated for 16 hours with undiluted mixture of MANDM4 and MANDM9 mAb culture supernatants (a), culture medium control (b), or mAb against fast myosin, followed by FITC-conjugated rabbit anti-mouse IgG (c).

To follow the maturation of normal and congenital DM fetal muscles, we analyzed the pattern of expression of MyHC isoforms in muscles from five CDM fetuses and five normal fetuses at an age between 16 and 35 weeks of development. Using specific antibodies against embryonic, fetal, and slow and fast MyHC isoforms, we measured the levels of the four isoforms in each muscle extract by Western blot (Figure 4) . Total MyHC in each muscle sample was first estimated by reactivity of the monoclonal antibody MF-20 that recognized all sarcomeric MyHC isoforms and protein loadings were adjusted to give approximately similar levels of total MyHC. Expression of embryonic and fetal MyHC disappeared or was greatly reduced in control muscles after 33 and 35 weeks of development, respectively. Embryonic MyHC is expressed during the early process of muscle differentiation, as is fetal MyHC, and both are replaced by fast or slow MyHC when the fibers become mature. Expression of embryonic MyHC in CDM muscles was still detected at 33 weeks of development. This would suggest that either new fibers are still being formed at this stage or the differentiation process is not yet complete. Expression of slow and fast MyHC increased from 16 to 35 weeks of development in control muscles, corresponding to the maturation of the fibers. In contrast, expression of slow MyHC was very low or almost absent in CDM muscles, whereas fast MyHC showed a similar pattern of expression to that observed in control muscles (Figure 4) . This would suggest that the maturation of the second generation of slow fibers is severely perturbed in the CDM fetal muscles.

View larger version (65K): [in this window] [in a new window]   Figure 4. Delayed maturation and absence of slow MyHC in CDM muscles. The pattern of expression of the different MyHC isoforms was determined in CDM muscles between 16 and 35 weeks of development. Expression of embryonic (emb), fetal, and fast and slow MyHC were analyzed by Western blot analysis using antibodies specific for each isoform and a pan-MyHC antibody. A dramatic reduction or total absence of slow MyHC and a persistence of developmental isoforms was observed in CDM muscles at term.

DMPK levels were measured in muscles from CDM and normal age-matched fetuses and results using the coil domain mAb, MANDM13, are shown in Figure 5A . DMPK was detected on Western blots and quantitated in Figure 5B , after applying a correction for protein loading as described in Materials and Methods. This experiment was repeated using MANDM1 to detect DMPK (data not shown) and the combined results are shown in Table 1 . DMPK in DM muscles was significantly reduced to 57% of control levels (P = 0.006, t-test). No clear correlation between CTG length and DMPK levels was observed in the muscle samples obtained from CDM fetuses with different CTG repeat lengths (between 1800 to 3700). In addition, all of the muscle samples used in this study had a dystrophin band at 427 kd (see lane 1 of the inset in Figure 5B ) and there was no consistent difference in dystrophin degradation between control and DM samples. This was performed to rule out the possibility that differential proteolysis might account for the reduced DMPK levels in DM muscle samples. Dystrophin is very sensitive to proteolysis and extracts without an intact 427-kd dystrophin band were rejected (see lane 2 of the inset in Figure 5B ). To confirm the result in Figure 5, A and B , we also compared the levels of DMPK in skeletal muscle cell cultures from CDM and age-matched normal fetal tissue. Using MANDM1 antibody, we observed that the level of DMPK in CDM skeletal cells was reduced to 53% of control values using the level of actin as internal control of equal loading (Figure 5C) . This experiment also rules out the possibility that differential postmortem proteolysis might account for the reduced DMPK levels.

View larger version (35K): [in this window] [in a new window]   Figure 5. Reduction of DMPK in muscles from CDM fetuses. DMPK levels in CDM and age-matched normal (control) muscles were determined by Western blot analysis using MANDM13 (A) and MANDM1 mAb (not shown, see Table 1 ). The histogram (B) represents the levels of DMPK in CDM and control muscles. Corrections for loading were applied to produce the histogram of control and DM levels. The number above each DM column is the number of CTG repeats in that fetus. The inset in B illustrates monitoring of proteolysis in muscle extracts by Western blotting with the anti-dystrophin mAb, MANDYS1; lane 1 contains some intact dystrophin whereas lane 2 has no 427-kd band and the extract would be rejected. DMPK levels were also reduced in CDM skeletal muscle cell cultures compared to age-matched controls (C). Actin is shown as a control for equal loading.

The expression of insulin receptor (InR) isoform A versus B in skeletal muscle of CDM and normal fetuses was determined by RT-PCR. Alternative splicing of the 36-nucleotide exon 11 of the -subunits of the InR pre-mRNA results in the expression of the two isoforms, A and B. InR-B that is found in the insulin-responsive tissues was expressed in skeletal muscle from normal fetuses whereas no expression of this isoform could be detected in the skeletal muscle from CDM fetuses (Figure 6) . Only the nonmuscle InR-A isoform was present in the CDM skeletal muscle. This effect on InR splicing seems to be due to changes in CUG-binding protein levels in DM1 nuclei that retain RNA transcripts with expanded CUG repeats.¹⁶

View larger version (29K): [in this window] [in a new window]   Figure 6. No B isoform of the insulin receptor in skeletal muscles from CDM fetuses. InR mRNA of A and B isoforms in skeletal muscle from CDM and nonaffected fetuses were determined by RT-PCR analysis. The exon 11 of the InR present only in the isoform B was not detectable in CDM skeletal muscle CDM.

	Discussion

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Characterization of human DMPK protein using antibodies has proven to be difficult and is still a source of confusion. In fact, different polyclonal antibodies raised against DMPK have detected proteins with various molecular weights from 45 to 85 kd. Our panel of monoclonal antibodies against DMPK eventually demonstrated that authentic human DMPK is a protein of 80 to 85 kd that is expressed almost exclusively in skeletal muscle and heart.²⁷ Analysis of DMPK expression during human muscle development using these highly specific monoclonal DMPK antibodies has now shown that the expression of the DMPK protein increased significantly in skeletal muscle between 9 and 16 weeks of development. This result is in good agreement with previous analysis that showed the presence of DMPK transcripts in skeletal muscle from a 13-week-old fetus.⁵³ Human skeletal muscle formation is a biphasic process with a first generation of myotubes forming in a relative synchronous manner between 8 and 10 weeks of development. Then, a second generation of fibers form progressively and asynchronously around these primary fibers between 11 and 18 weeks of development.⁵⁴ The large increase in accumulation of DMPK protein correlates with the formation of this second generation of muscle fibers and the major period of muscle formation. After 20 weeks, no new fibers are formed and the muscle fibers undergo a process of maturation. DMPK levels remained high during this time (Figure 2) . This is consistent with evidence that DMPK transcription in muscle cells is under the control of muscle-specific regulatory elements located in the promoter and the first intron of the DMPK gene.⁵⁵ The increase in DMPK expression during in vitro myogenic differentiation (Figure 1) is in good agreement with the appearance of DMPK protein during muscle formation and the persistence of high levels of DMPK during muscle growth.

To study the effects of CTG repeat expansion on DMPK levels, fetal muscle samples from DM1 patients are difficult to obtain. We were fortunate, however, in finding samples with age-matched controls at a number of different fetal ages. We would have preferred to have multiple samples at each fetal stage to determine the statistical significance of differences between CDM and control at each stage. Although this was not possible, a paired t-test on all of the paired samples showed a highly significant reduction in DMPK protein levels overall (Table 1) . We restricted our analysis to CDM fetuses with very large CTG repeat expansions (>1800 CTG). If large CTG expansions cause complete nuclear retention of mutated transcripts and/or altered its transcription, cytoplasmic DMPK mRNA would still only be reduced by 50% (because only one of two DMPK alleles is affected). However, feedback regulation of DMPK levels might up-regulate DMPK protein production. Our studies suggest that this up-regulation (by increased translation or decreased turnover) does not occur in fetal muscle because the level of DMPK in CDM fetal muscles is 57% of controls and a similar reduction in DMPK levels was also observed in cultured muscle cells. These results are entirely predictable from what we understand to be the mechanism for the reduction. An earlier study reported only a slight change in the level of DMPK between muscle samples from CDM patients and age-matched controls,⁵⁶ whereas in our study we have found a much larger and significant decrease in the level of DMPK in muscles of CDM fetuses. This can be explained by the fact that our samples differed significantly from those of Narang and colleagues.⁵⁶ We have used muscle samples from fetuses instead of 2- to 6-day-old infants and, more importantly perhaps, our samples had much larger CTG repeat expansions (range of 1800 to 3700 instead of 520 to 910). We have used a very specific DMPK monoclonal antibody in this study and we were also able to demonstrate a significant decrease (53%) of DMPK levels in CDM muscle cell cultures. Similar reductions in DMPK levels were also found in DM1 muscle cells.⁵⁷ This is consistent with an almost complete nuclear retention of the mutant DMPK transcripts since we have shown previously that foci of mutated DMPK RNA were present in these CDM muscle cells in culture.⁵⁸ However, there is little evidence that reduced DMPK levels have a deleterious effect in skeletal muscle. Heterozygotes of DMPK knockout mice, in which a similar reduction in DMPK levels can be expected, show no skeletal muscle phenotype, the only effects being on cardiac conduction. In contrast, DM features were observed in transgenic mice expressing expanded CUG repeats, whether or not the repeats were within DMPK transcripts.^23,24

Previous analysis of muscle from CDM patients showed that muscle fibers were less mature in fetuses and that skeletal muscle maturation was impaired in preterm infants.⁹ This delay in maturation of the skeletal muscle was characterized by the abnormal presence of myotubes, smaller fascicles of muscle fibers, thinner or atrophic myofibers, and delayed differentiation of the second generation of fibers.¹⁰ The maturation of skeletal muscle can be followed by analyzing the pattern of expression of the MyHC isoforms because a progressive switch from embryonic and fetal to slow and fast MyHCs occurs during this process. In the present study performed on muscle biopsies from unaffected and CDM-affected fetuses between 16 and 35 weeks of development, we showed a delay of muscle maturation in CDM fetuses with a persistence of the embryonic and fetal MyHCs after 30 weeks of development and almost complete absence of the slow MyHC. This abnormal phenotype is probably caused in some way by the large CUG repeats in retained nuclear transcripts, because neither the DMPK nor the Six5 knockout mice show any change in muscle formation or fiber-type expression.^20,21 It has been suggested that the nuclear accumulation of expanded CUG-repeat RNA leads to aberrant RNA processing by altering the levels of the CUG-BP proteins. In this study, we have demonstrated an abnormal regulation of insulin receptor RNA splicing in the muscles of our CDM fetuses (Figure 6) and this, or a similar aberrant splicing event, could be involved in the delayed fiber maturation in CDM. We have demonstrated previously that large CTG repeats altered the behavior of CDM satellite cells in vitro.⁵⁸ Both the proliferative capacity and the myogenic differentiation of CDM myogenic precursor cells were found to be defective. The maturation and growth of the muscle are associated with the addition of myonuclei resulting from proliferation and fusion of activated satellite cells. Because both are reduced in CDM muscle cells in vitro, we suggested that these anomalies could be responsible at least in part for the delay in maturation and muscle atrophy that is observed in the CDM fetuses. However, we cannot exclude the possibility that reduced levels of DMPK protein might increase disease severity. In particular, it has been suggested that slow fibers might be more DMPK-dependent than fast fibers because DMPK protein is exclusively expressed in slow fibers.^43-45 However, by Western blot analysis of isolated slow and fast single fibers, we have demonstrated that DMPK protein is expressed in both fiber types (Figure 3A) . In addition, immunohistochemical analysis on frozen tissue sections, using our DMPK monoclonal antibodies, did not reveal any differential staining between fast and slow fibers (Figure 3B) . This result is supported by a recent study that found DMPK RNA transcripts in both fast and slow fibers and no reduction in slow fibers from adult DM1 patients.⁵⁹ Maturation of slow fibers depends on innervation,⁶⁰ so any delay or defect in the function of the neuromuscular junction during the fetal period could also contribute to the almost complete absence of the slow MyHC.

The severe congenital form of DM1 associated with large CTG repeats showed delayed muscle development. The importance in DM1 pathogenesis of a gain of function involving expanded CUG repeats in DMPK transcripts is now well documented. This mechanism rather than DMPK reduction seems to be implicated in the abnormal CDM muscle phenotype.

	Acknowledgements

 
We thank Dr. J. P. Barbet and the Tissue Bank of the Association Française contre les Myopathies for providing some of the tissue samples.

	Footnotes

 
Address reprint requests to Prof. G. E. Morris, D. Phil., Biochemistry Group, North East Wales Institute, Mold Road, Wrexham, LL11 2AW, UK. E-mail: morrisge{at}newi.ac.uk

Supported by the Muscular Dystrophy Campaign, Association Française contre les Myopathies, CNRS, and the European Community (contract QLK6-1999-02034).

D. F. and L. T. L. contributed equally to this work.

Accepted for publication November 15, 2002.

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