Increased expression of utrophin in a slow vs. a fast muscle involves posttranscriptional events

Anthony O. Gramolini, Guy Bélanger, Jennifer M. Thompson, Joe V. Chakkalakal, and Bernard J. Jasmin

Department of Cellular and Molecular Medicine, Faculty of Medicine, and Centre for Neuromuscular Disease, University of Ottawa, Ottawa, Ontario, Canada K1H 8M5


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In addition to showing differences in the levels of contractile proteins and metabolic enzymes, fast and slow muscles also differ in their expression profile of structural and synaptic proteins. Because utrophin is a structural protein expressed at the neuromuscular junction, we hypothesize that its expression may be different between fast and slow muscles. Western blots showed that, compared with fast extensor digitorum longus (EDL) muscles, slow soleus muscles contain significantly more utrophin. Quantitative RT-PCR revealed that this difference is accompanied by a parallel increase in the expression of utrophin transcripts. Interestingly, the higher levels of utrophin and its mRNA appear to occur in extrasynaptic regions of muscle fibers as shown by immunofluorescence and in situ hybridization experiments. Furthermore, nuclear run-on assays showed that the rate of transcription of the utrophin gene was nearly identical between EDL and soleus muscles, indicating that increased mRNA stability accounts for the higher levels of utrophin in slow muscles. Direct plasmid injections of reporter gene constructs showed that cis-acting elements contained within the utrophin 3'-untranslated region (3'-UTR) confer greater stability to chimeric LacZ transcripts in soleus muscles. Finally, we observed a clear difference between EDL and soleus muscles in the abundance of RNA-binding proteins interacting with the utrophin 3'-UTR. Together, these findings highlight the contribution of posttranscriptional events in regulating the expression of utrophin in muscle.

Duchenne muscular dystrophy; synaptic proteins; messenger ribonucleic acid stability; ribonucleic acid-binding proteins; neuromuscular junction


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

DUCHENNE MUSCULAR DYSTROPHY (DMD) is the most prevalent inherited neuromuscular disorder, affecting 1 of every 3,500 male births (15). The disease is characterized by repetitive cycles of muscle degeneration-regeneration, with fast-contracting fibers being preferentially affected (56). Eventually, the muscle's regenerative capacity fails, and the muscle mass is progressively replaced by adipose and connective tissues. The disease is extremely severe because children become wheelchair bound by early adolescence and death usually occurs in their second or third decade of life. DMD results from mutations/deletions in the X-linked dystrophin gene, which prevents production of a large cytoskeletal protein of the spectrin superfamily termed dystrophin (37, 58). Several approaches have been envisaged to counteract the deleterious effects of this progressive disease, including, for example, gene therapy and cell transfer. An alternative strategy consists in using a protein normally expressed in dystrophic muscle which, once expressed at appropriate levels and at the correct subcellular location, could functionally compensate for the lack of dystrophin. A good candidate for such a role is utrophin, primarily because of its high degree of sequence identity with dystrophin (53). In addition, several recent studies using transgenic mouse model systems have clearly shown the ability of utrophin to functionally compensate for the lack of dystrophin (for example, see Ref. 54).

In contrast to dystrophin, which is found along the entire length of normal muscle fibers (37, 58), utrophin accumulates preferentially at the postsynaptic membrane of the neuromuscular junction in both normal and DMD muscles (4, 24). In this context, it has been shown that local transcriptional activation of the utrophin gene in myonuclei located within the postsynaptic sarcoplasm accounts, at least partially, for the preferential expression of utrophin at the neuromuscular junction (18, 20, 21, 31, 55). Interestingly however, there are several cases in which expression of utrophin has been shown to extend well into extrasynaptic regions of muscle fibers. For example, utrophin is known to be present outside synaptic regions in small or regenerating muscle fibers of DMD patients (26, 30, 40). Moreover, increased levels of utrophin have been reported along the length of developing fibers in both embryonic and neonatal muscles (30, 32, 42, 50). Under specific conditions, therefore, utrophin presents a more homogeneous distribution along muscle fibers, indicating that additional regulatory mechanisms are likely involved in controlling the overall levels and localization of utrophin in skeletal muscle.

It is well established that adult fast and slow skeletal muscles differ markedly in their physiological characteristics as a result of pronounced differences in the expression of numerous contractile proteins and metabolic enzymes (for review, see Refs. 44, 48, and 51). In addition to these well-known differences, recent studies have also demonstrated that fast and slow muscles contain varying amounts of structural proteins. In particular, slow muscles express significantly more spectrin (38, 57) and dystrophin (11, 28). Interestingly, there is also evidence showing that differences between fast and slow muscles extend even to the pattern of expression of transcripts encoding synaptic proteins. Indeed, mRNAs encoding acetylcholinesterase (AChE) (39), acetylcholine receptor (AChR) subunits (34), and ColQ (33) are found expressed at significant levels in extrasynaptic regions of slow muscle fibers. Because utrophin is a structural protein preferentially expressed at the neuromuscular junction, we hypothesize in the present study that the expression and localization of utrophin is different between fast and slow muscles. Because our results indicated that slow muscles do, in fact, contain more utrophin than fast muscles, we also became interested in identifying the molecular mechanisms that account for this difference.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Animal care. Control C57BL/6 mice were obtained from Charles River Laboratories (St. Constant, Quebec, Canada) and housed in the University of Ottawa Animal Care Facility. All surgical procedures were performed in accordance with the strict guidelines established by the Canadian Council on Animal Care. For most of these studies, the slow soleus and fast extensor digitorum longus (EDL) muscles were used. In some cases, the lateral and medial gastrocnemius muscles were also excised. Because we were also interested in determining utrophin levels in the fast and slow regions of a given muscle, these muscles were further dissected into their red and white compartments. After excision, muscles were either flash frozen in liquid nitrogen or embedded in optimum cutting temperature compound and frozen in melting isopentane precooled with liquid nitrogen.

Characterization of reporter gene constructs and direct gene transfer. The full-length utrophin 3'-untranslated region (3'-UTR) was generated by RT-PCR using RNA isolated from C2C12 cells and primers designed against the mouse utrophin sequence (25). The PCR product was sequenced and compared with sequences available in data banks (Gramolini, Bélanger, and Jasmin, unpublished observations). The full-length utrophin 3'-UTR (1,995 nt) was subsequently inserted downstream of the reporter gene LacZ in the pCMV SPORT beta -galactosidase expression vector (GIBCO BRL, Burlington, Ontario, Canada) in the forward or reverse (used as a control) orientation.

Direct gene transfer was performed as described in detail elsewhere (10, 20, 21) with the exception that soleus and EDL muscles were injected bilaterally with plasmid DNA instead of tibialis anterior muscles. Briefly, a small incision was made on the lateral side of both hindlimbs while the mice were anesthetized. The soleus and EDL muscles were carefully isolated and injected with 25 µl of the appropriate plasmid cDNAs (2-4 µg/µl) under a dissecting microscope. The wounds were then closed using autoclips, and the animals were returned to their cage. Seven to ten days later, injected muscles were excised and immediately frozen in liquid nitrogen. The injection solution contained, in addition to the appropriate reporter constructs (see above), a constitutively expressed chloramphenicol acetyltransferase (CAT) plasmid (Promega), which was coinjected to monitor the efficiency of transduction. In all these experiments, plasmid DNA was prepared using the Qiagen Mega-prep and/or Mini-prep procedures (Chatsworth, CA).

RNA extraction and RT-PCR. Total RNA was extracted from muscles using TriPure as recommended by the manufacturer (Boehringer Mannheim, Indianapolis, IN). Briefly, soleus and EDL muscles were homogenized in 1 ml of TriPure reagent using a Polytron. After chloroform addition, the homogenates were centrifuged at 12,000 g for 10 min at 4°C. The resulting aqueous phase was transferred to a fresh microfuge tube, and 0.5 ml of ice-cold isopropanol was added. The RNA pellets were isolated by centrifugation at 12,000 g for 15 min and thoroughly washed with 75% ethanol. The pellets were finally resuspended in 20 µl of RNase-free water and stored at -20°C. Before RT-PCR, RNA samples were treated with RQ DNase I (Promega, Madison, WI) at 37°C for 90 min and heated at 65°C for 20 min to terminate the reaction

Quantitative RT-PCR was used to strictly determine the relative abundance of utrophin transcripts in fast vs. slow muscles. This assay was performed as described in detail previously (18, 20, 29, 39). Reverse transcription was carried out at 42°C using 100 ng of total RNA and random hexamers. After 45 min, the RT mixtures were heated to 99°C for 5 min to terminate the reaction. Primers that selectively amplified utrophin (20, 29) and S12 rRNA (6, 16) were designed on the basis of available sequences. These primers amplified a 548-bp and a 368-bp fragment from the mouse utrophin and S12 cDNAs, respectively. To examine expression of utrophin A and B mRNAs, primers against the unique 5'-UTR of these transcripts were designed on the basis of available sequences (8). These primers amplify a 530-bp and a 101-bp fragment in the utrophin A and B transcripts, respectively. cDNAs encoding beta -galactosidase were amplified using primers that generate a 506-bp fragment (5). PCR amplification was performed in a DNA thermal cycler (Perkin Elmer Cetus, Norwalk, CT). Each cycle of amplification consisted of denaturation at 94°C for 1 min, primer annealing at 65°C for 1 min, and extension at 72°C for 1 min. In these assays, negative controls consisted of reverse transcription mixtures in which total RNA was replaced with RNase-free water.

The PCR products were visualized on 1% agarose gel containing ethidium bromide. The 100-bp molecular weight markers (MBI Fermentas, Flamborough, Ontario, Canada, and GIBCO BRL) were used to estimate the size of the PCR products. For quantitative experiments, PCR products were separated and visualized on agarose gels containing the fluorescent dye Vistra Green (Amersham, Arlington Heights, IL) (20). The labeling intensity of the PCR product, which is linearly related to the amount of DNA, was subsequently quantitated using a Storm PhosphorImager and the accompanying Imagequant software (Molecular Dynamics, Sunnyvale, CA). Values obtained for utrophin were standardized relative to the corresponding level of rRNA in the same sample. All RT-PCR measurements aimed at determining the relative abundance of selected transcripts were performed during the linear range of amplification (see, for example, Refs. 9 and 39). Typically, the cycle numbers were 26 to 30. RT-PCR conditions (primer concentrations, input RNA, choice of RT primer, cycling conditions) were initially optimized, and these were identical for all samples. Appropriate precautions were taken to avoid contamination and RNA degradation. All samples as well as negative controls were prepared using common master mixes containing the same RT and PCR reagents, and they were always run in parallel. In all experiments, PCR products were never detected for the negative controls.

In situ hybridization. Longitudinal serial cryostat sections (12 µm) from EDL and soleus muscles were cut in a cryostat and placed on Superfrost microscope slides (VWR Canlab). Slides were first processed for AChE histochemistry, and the regions containing neuromuscular junctions were photographed. The sections were then subjected to in situ hybridization using synthetic oligonucleotides for detection of utrophin transcripts as described (21). The in situ hybridization experiments were performed using two antisense oligonucleotides complementary to mouse utrophin transcripts (21).

Quantitative analysis of in situ hybridization labeling was performed using an image analysis system equipped with Northern Eclipse software (Empix Imaging, Mississauga, Ontario, Canada). Briefly, labeling density in synaptic vs. extrasynaptic regions was determined by measuring the pixel intensity within circular fields of similar sizes in muscle fibers and by subtracting background values determined as the signal seen in regions external to the fibers (21, 23, 41). For these analyses, both EDL and soleus muscle sections were placed on the same slide and processed for in situ hybridization simultaneously. Three separate experiments were performed using a minimum of ten muscle sections per condition. A minimum of four measurements were performed on each cryostat section.

Immunoblotting. Muscles were homogenized using a Polytron in Tris · HCl, pH 8.0 (1% sodium deoxycholate, 5% SDS, 0.5% Triton X-100, 1 mM phenylmethylsulfonyl fluoride (PMSF), 5 mM iodoacetamide, 2 mg/ml aprotinin, 100 mM Tris · HCl, 140 mM NaCl, and 0.025% NaN3) and subjected to immunoblotting as described (20). For some of these experiments, rat soleus and EDL muscles were also used with similar results. Briefly, equivalent amounts of proteins (up to 200 µg) were separated on a 6% polyacrylamide gel and electroblotted onto a polyvinylidene difluoride membrane (Sigma, St. Louis, MO). After transfer, the membranes were incubated with a monoclonal antibody directed against utrophin (Novocastra Laboratories, Newcastle upon Tyne, UK) and revealed using a commercially available chemifluorescence kit (NEN Life Sciences, Boston, MA). To ensure that equivalent amounts of proteins were loaded for each sample, membranes were also stained with Ponceau S (Sigma).

Immunofluorescence. Immunofluorescence experiments were performed on longitudinal serial cryostat sections (12 µm) of EDL and soleus muscles placed on the same slide. Utrophin immunoreactivity was detected using a monoclonal anti-utrophin antibody (Novocastra) and a goat anti-mouse Cy3-conjugated secondary antibody (Jackson Laboratories; Bio/Can Scientific, Mississauga, Ontario, Canada). Fluorescein isothiocyanate-conjugated alpha -bungarotoxin (Molecular Probes, Eugene OR) was used to label AChR at the neuromuscular junctions.

Isolation of nuclei and in vitro transcription assays. Nuclei were isolated from soleus and EDL muscles, and in vitro transcription assays were performed as described (5, 9, 22). Briefly, pools of 12-14 muscles were first washed with PBS and homogenized with a Dounce homogenizer in a solution containing 10% sucrose, 60 mM KCl, 15 mM NaCl, 15 mM HEPES, 0.5 mM EGTA, 2 mM EDTA, 0.1 mM spermine, 0.5 mM spermidine, 0.5 mM dithiothreitol (DTT), and 1 µM PMSF. The nuclei were then isolated by centrifugation. They were resuspended in a solution containing 50% glycerol, 20 mM Tris, pH 7.9, 75 mM NaCl, 0.5 mM EDTA, 0.5 mM DTT, 1 µM PMSF, and 10 U/µl RNase inhibitor and subjected to in vitro transcription by adding 200 µCi of [alpha -32P]UTP (Amersham) to label nascent transcripts for 30 min at 27°C. After DNase I digestion and protein denaturation, radiolabeled RNA was extracted using TriPure (see above) and hybridized to Protran nitrocellulose membranes (Schleicher and Schuell, Keene, NH) containing 10 µg of immobilized genomic DNA and cDNAs encoding utrophin (41). After hybridization, the membranes were washed thoroughly (1× standard saline citrate, 0.1% SDS) at 42°C and subjected to autoradiography. The signal intensities were quantitated using a Storm PhosphorImager, and the intensity of the utrophin signals were standardized according to the corresponding signal seen with genomic DNA.

In vitro transcription and ultraviolet crosslinking. The utrophin 3'-UTR was subcloned into pCR2.1-TOPO vector (Invitrogen). In vitro RNA-binding studies were performed as previously described (2, 5). Briefly, 32P-labeled sense RNA was generated using the T7 polymerase. Labeled RNA (~5 × 104 counts/min) was then incubated with 100 µg of total protein extracts isolated from the soleus or EDL muscles. To obtain protein extracts, muscles were homogenized using a Polytron in 10× (mass/volume) of a buffer containing 0.3 M sucrose, 60 mM NaCl, 15 mM Tris (pH 8.0), 10 mM EDTA, 0.1 mM beta -mercaptoethanol, 0.01 mM PMSF, 0.01 mM benzamidine, 1 µg leupeptin, 10 µg pepstatin A, and 1 µg aprotinin (2). The extracts were centrifuged at 14,000 g for 15 min at 4°C. The supernatants were then removed, and the protein concentration was determined using the bicinchoninic acid protein assay kit (Pierce Laboratories, Rockford, IL). Samples were stored at -80°C until further analysis.

RNA protein complexes were crosslinked by irradiation for 30 min at 4°C at 1 cm from an ultraviolet (UV) lamp (3,000 mW/cm2) in the presence of RNase inhibitors. Free RNA was then digested with RNase A and T1, and the UV-cross-linked products were analyzed by 10% SDS-polyacrylamide gel electrophoresis and by autoradiography. For competition experiments, a 10-fold molar excess of unlabeled probe was incubated 15 min before the radioactive probe was added.

Statistical analysis. Paired Student's t-tests were performed to evaluate the differences in utrophin expression between EDL and soleus muscles as well as between synaptic and extrasynaptic regions of individual muscle fibers. The level of significance was set at P < 0.05. Data are expressed as means ± SE throughout.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Slow muscles contain more utrophin. To determine whether utrophin expression varies between different muscle types, we first examined the levels of utrophin in the slow soleus as well as fast EDL muscles. As shown in Fig. 1, immunoblotting experiments revealed that the levels of utrophin are approximately three- to fourfold higher (P < 0.05) in soleus muscles compared with the levels seen in EDL muscles. To ascertain that this difference between these muscles could be extended to other muscles and that it was indeed related to muscle type and not to muscle function, i.e., ankle extensors vs. ankle flexors, we also examined the relative levels of utrophin in the red and white portions of the lateral and medial gastrocnemius muscles. As illustrated in Fig. 1C, the red portions of the medial and lateral gastrocnemius muscle contain considerably more utrophin compared with the white, faster-contracting adjacent regions (Fig. 1C).


View larger version (37K):
[in this window]
[in a new window]
 
Fig. 1.   Expression of utrophin in fast vs. slow muscles. A: an example of a Western blot showing the presence of utrophin in extract from extensor digitorum longus (EDL; fast) and soleus (SOL; slow) muscles. Coomassie blue staining of the gel confirmed equal loading (not shown). B: quantitation indicating that, compared with fast muscles, slow muscles contain approximately 3- to 4-fold more utrophin. Shown are results obtained with 4 independent experiments. * Significant difference between the 2 groups (P < 0.05). C: examples of Western blots showing the levels of utrophin in the red (R) and white (W) portions of the medial and lateral gastrocnemius muscles. Similar to the findings with soleus muscles, the red portions of the gastrocnemius muscle contain more utrophin. Shown are examples obtained with a minimum of 5 muscles.

We next performed immunofluorescence experiments to confirm these observations using light microscopy and to gain insights into the distribution of utrophin in slow vs. fast muscles. In these experiments, we observed, as expected, an accumulation of utrophin at the neuromuscular junctions from both soleus and EDL muscles (Fig. 2). However, we also determined that utrophin is present at low levels throughout the sarcolemma of soleus muscle fibers. These findings indicate, therefore, that the greater amount of utrophin detected in soleus muscles results from an increase in its expression in extrasynaptic regions of muscle fibers.


View larger version (94K):
[in this window]
[in a new window]
 
Fig. 2.   Localization of utrophin in EDL vs. soleus muscles. Shown are representative examples of photomicrographs of EDL (A and B) and soleus (C and D) muscles processed for double-fluorescence experiments using fluorescein-conjugated alpha -bungarotoxin, which recognizes acetylcholine receptors (AChR) located at the neuromuscular junctions (A and C), and an antibody against utrophin (B and D). In addition to the expected junctional accumulation, note the expression of utrophin at the sarcolemma in extrasynaptic compartments of soleus muscle fibers. Bar, 50 µm.

Increased expression of utrophin transcripts in soleus vs. EDL muscles. To determine whether these differences in the abundance of utrophin were paralleled by changes in the expression of its mRNA, we next performed a series of RT-PCR analyses and in situ hybridization experiments. For these experiments, we chose to focus on the soleus and EDL as representative of slow and fast muscles, respectively, since these two muscles are of similar size. In addition, they are easily accessible, thereby increasing the reliability of the dissection procedure and injection experiments (see below). Consistent with the immunoblotting results, we observed that the levels of utrophin mRNAs were higher in soleus vs. EDL muscles (Fig. 3A). Quantitative measurements of the relative abundance of utrophin transcripts revealed that, compared with their fast-twitch counterparts, soleus muscles contain significantly more (P < 0.05) utrophin mRNAs (Fig. 3B). Because skeletal muscles express two distinct utrophin transcripts termed A and B, which arise from different promoters (8), we also determined whether the expression of both transcripts was similarly increased in slow muscles. To this end, we selectively amplified the utrophin A or B mRNA by PCR using primers designed against the unique 5'-UTR of these transcripts (8). These experiments showed that both utrophin A and B transcripts were more abundant in the slow soleus muscle (data not shown).


View larger version (22K):
[in this window]
[in a new window]
 
Fig. 3.   Expression of utrophin mRNA in EDL vs. soleus muscles. A: an ethidium bromide-stained agarose gel showing RT-PCR products for utrophin and S12 rRNA (internal standard) obtained from the mouse soleus (slow) and EDL (fast) muscles. The negative control lane is marked with a minus sign. Left lane: the 100-bp molecular mass marker (MBI Fermentas). B: compared with their fast-twitch counterparts, slow muscles contain ~4-fold more utrophin transcripts. Shown are results obtained from 6 independent experiments. * Significant difference between the 2 groups (P < 0.05).

To further characterize the increase in utrophin mRNA levels in soleus muscles and to confirm the RT-PCR data, we also performed in situ hybridization experiments aimed at localizing utrophin transcripts along single soleus and EDL muscle fibers. Because our RT-PCR data showed that the utrophin A and B transcripts were similarly increased in soleus muscles, we used synthetic oligonucleotides that recognize both mRNA species (21). Although utrophin transcripts are enriched at the neuromuscular junction in both muscle types (Fig. 4; see also Refs. 21 and 55), we observed a clear increase in the abundance of utrophin transcripts in extrajunctional regions of soleus fibers. Quantitative analyses revealed, in agreement with our immunofluorescence data, that the synaptic enrichment of utrophin mRNAs was similar in fast and slow muscles, whereas in extrajunctional compartments utrophin transcripts were approximately threefold (P < 0.05) more abundant in soleus muscle fibers (Fig. 4C).


View larger version (52K):
[in this window]
[in a new window]
 
Fig. 4.   Localization of utrophin mRNA in EDL and soleus muscles. A and B show examples of cryostat sections from the EDL (fast) and soleus (slow) muscles, respectively, processed for in situ hybridization using synthetic oligonucleotides that recognize utrophin transcripts. Each section shows a synaptic region (accumulation of grains; see arrows) and extrasynaptic compartments. Bar, 100 µm. As observed by RT-PCR, the slow soleus muscle (C) contains more utrophin mRNAs. This experiment also indicates that the greater abundance of utrophin transcripts in slow muscle is due to an enrichment in extrasynaptic regions. * Significant difference between the 2 groups (P < 0.05).

The utrophin gene appears transcribed at a similar rate in EDL and soleus muscles. To determine whether the increased utrophin mRNA and protein expression seen in soleus muscles results from a greater transcriptional activity of the utrophin gene, we performed run-on transcription assays using nuclei isolated from EDL and soleus muscles. In these experiments, we observed within the sensitivity of this approach that the rate of transcription for the utrophin gene appeared nearly identical in both muscle types (Fig. 5A). Densitometric analysis of these results confirmed that the transcriptional activity of the utrophin gene in soleus muscles was not significantly different (P > 0.05) from the activity observed in EDL muscle (Fig. 5B). As a positive control for these assays, we observed that transcription of the gene encoding the glycolytic enzyme glyceraldehyde-3-phosphate dehydrogenase was, as expected, higher in EDL samples (not shown). Furthermore, in parallel experiments using the same approach, we recently described significant changes in gene transcription (see, for example, Ref. 1). Therefore, our findings indicate that enhanced stability of existing transcripts likely accounts for the higher levels of utrophin mRNAs in soleus muscles (see Fig. 3).


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 5.   Transcriptional activity of the utrophin gene in EDL vs. soleus muscles. A: examples of a nuclear run-on assay performed with nuclei isolated from a pool of EDL (fast) and soleus (slow) muscles to detect nascent utrophin transcripts. B: the quantitative analysis of 3 independent experiments showing that the transcriptional activity of the utrophin gene is nearly identical in slow and fast muscles (P > 0.05), thereby indicating that the stability of existing transcripts likely accounts for the differences in utrophin mRNA levels in these 2 muscle types.

The utrophin 3'-UTR regulates expression of a chimeric reporter construct in soleus muscles. On the basis of these observations, we next performed a series of experiments aimed at examining the importance of posttranscriptional mechanisms in the regulation of utrophin in fast vs. slow muscles. We thus performed direct plasmid injection experiments using constitutively expressed LacZ reporter constructs engineered to contain the utrophin full-length 3'-UTR, which is common to both A and B transcripts. Soleus and EDL muscles were injected with this construct and with a CAT plasmid used to monitor the efficiency of transduction. Seven to ten days later, expression of LacZ transcripts was examined by RT-PCR. As show in Fig. 6A, levels of the reporter transcript were clearly higher in soleus muscles compared with the levels seen in the EDL. In fact, quantitative analysis (n = 8) revealed that LacZ expression, normalized to CAT mRNA levels, was approximately two- to threefold higher in slow muscles. By contrast, we failed to detect any significant difference in the expression of LacZ transcripts in soleus and EDL muscles injected with a reporter construct containing the utrophin 3'-UTR in the reverse orientation and used, in this case, as a control (Fig. 6B).


View larger version (26K):
[in this window]
[in a new window]
 
Fig. 6.   Expression of a reporter gene construct in EDL and soleus muscles. A: a typical example (n = 8) of an ethidium bromide-stained agarose gel of RT-PCR products corresponding to chimeric LacZ transcripts expressed in EDL (fast) and soleus (slow) muscles after direct injection of a reporter construct containing the utrophin 3'-untranslated region (3'-UTR) in the forward orientation. In agreement with our hypothesis, expression of the reporter gene is higher in the slow soleus muscle, indicating that elements in the utrophin 3'-UTR enhance the stability of reporter transcripts in slow muscle. B: the same as A except that in this case the LacZ reporter construct contained the utrophin 3'-UTR in the reverse orientation. As expected with this control experiment, note the lack of difference in the levels of LacZ transcripts between fast and slow muscles.

On the basis of these results, we expected that the pattern of RNA-protein interactions would be different between fast and slow muscles. To examine this, we performed UV-cross-linking experiments by incubating protein extracts from soleus or EDL muscles, with in vitro-transcribed 32P-labeled RNA corresponding to the utrophin 3'-UTR. Using this approach, we determined the number of possible cytoplasmic factors that could bind to the 3'-UTR as well as their molecular mass. In protein extracts from soleus muscles, we readily detected two protein complexes (arrows in Fig. 7). The presence of these complexes was completely abolished by preincubation of the protein extracts with an excess of cold unlabeled probe. In agreement with our hypothesis and with the results presented above, we also observed an increase in the expression of these RNA-binding proteins in fast muscles because the intensity of the most abundant complex was clearly greater in EDL muscles.


View larger version (51K):
[in this window]
[in a new window]
 
Fig. 7.   RNA-protein interactions in EDL vs. soleus muscles: an ultraviolet (UV)-cross-linking experiment using the utrophin 3'-UTR and cytoplasmic extract from EDL (fast) and soleus (slow) muscles. These experiments were performed by incubating 100 µg of muscle extract with in vitro-transcribed 32P-labeled RNA corresponding to the utrophin 3'-UTR in the absence (-) or presence (+) of an excess of cold (unlabeled) probe. The bands at ~42 and 90 kDa (arrows) are more abundant in the fast EDL muscle, and these 2 bands were specifically competed by the excess of cold probe.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Although our recent studies (18, 20, 21), as well as those of others (31, 55), have shown that transcriptional regulatory mechanisms account, at least partially, for the preferential synaptic accumulation of utrophin, it is nonetheless possible to envisage that posttranscriptional events are also involved in the regulation of utrophin in muscle fibers. In this context, we have shown that distinct regions in the 3'-UTR control the targeting and stability of utrophin mRNAs in cultured muscle cells (Gramolini, Bélanger, and Jasmin, unpublished observations). Additionally, a recent study has nicely highlighted the contribution of posttranscriptional mechanisms in the development and maintenance of neuromuscular junctions in Drosophila (49). In agreement with these findings, we now show that utrophin is more abundant in soleus vs. EDL muscles as a result of an increase in the stability of its mRNAs, which appears mediated by cis-acting elements in the 3'-UTR and by the availability of RNA-binding proteins. Together, these data show, therefore, that in addition to transcriptional regulation, posttranscriptional events also play a key role in controlling the expression and localization of utrophin in skeletal muscle.

In our experiments, we also observed that the red portion of both the medial and lateral gastrocnemius muscle contains more utrophin compared with their respective adjacent white compartment. Therefore, it appears likely that the importance of posttranscriptional mechanisms generally operates in slow muscle fibers, thereby regulating the abundance and localization of utrophin. Even though there appears, on the basis of these data, to be a good correlation between muscle type, i.e., fast vs. slow, and utrophin expression, additional work remains to be done to refine this relationship by correlating, for example, utrophin levels with the expression of proteins known to be excellent markers of the contractile speed or fatigue characteristics of individual muscle fibers.

In general, it has become evident that, for a number of genes, transcriptional regulation cannot solely account for the observed changes in mRNA levels known to occur under a variety of conditions. Thus the importance of mRNA stability is becoming increasingly recognized as a key regulatory step in the control of gene expression (12, 45, 46). In this context, it is known that the half-life of distinct mRNAs within the same cell can vary by more than one order of magnitude, ranging from minutes for relatively labile transcripts to hours or even days in the case of unusually stable mRNAs. More interesting is the fact that these half-lives can be specifically altered according to environmental and/or intracellular influences (35, 45). Indeed, there are numerous examples using cultured cells that have shown that hormones, growth factors, and ionic concentrations can affect the stability of presynthesized mRNAs, thereby leading to rapid and dramatic changes in gene expression (46).

In skeletal muscle cells, it has also become clear that posttranscriptional mechanisms are indeed important in dictating the abundance of specific mRNAs (for example, see Refs. 52 and 59). Interestingly, there are even a few cases where the importance of posttranscriptional events has been implied in the expression of genes encoding synaptic proteins in muscle. In particular, a series of studies have shown that the increased expression of AChE mRNA that occurs during differentiation of myogenic cells in culture results in part from the enhanced stability of existing transcripts (3, 17). Consistent with these observations, we have recently determined that the ~10-fold reduction in AChE mRNA levels that occurs in denervated rat muscles results from a reduction in the half-life of existing transcripts with little change in the rate of transcription of the AChE gene (5). Similarly, the contribution of mRNA stability has to be taken into account in the response of AChR subunit genes to denervation since the ~5- to 10-fold increase in transcription, known to occur in denervated muscles, is transient and cannot fully account for the much greater increases seen in the abundance of AChR transcripts (7, 14). Finally, in a collaborative effort, we have shown that the synaptic accumulation of alpha -dystrobrevin 1 transcripts in muscle fibers likely involves posttranscriptional mechanisms as opposed to a local transcriptional activation of the dystrobrevin gene in myonuclei located in the postsynaptic sarcoplasm (41). Taken together with our present findings, results of these studies indicate therefore that, in addition to transcriptional regulation (7, 14, 47), posttranscriptional mechanisms represent key events for controlling the expression and localization of synaptic proteins in muscle fibers.

Although there is now more information illustrating the importance of mRNA stability in the control of gene expression, there are relatively few examples where the precise molecular mechanisms have been clearly defined. For instance, several unstable mRNAs, including c-fos and granulocyte-macrophage colony-stimulating factor, contain adenosine-uridine-rich elements (AURE) in their 3'-UTR that regulate their turnover through interactions with members of the AURE-binding protein family. Additionally, mRNAs encoding the transferrin receptor contain iron-responsive elements that bind a cytosolic protein referred to as IRP, which stabilizes these transcripts by preventing their degradation during iron deficiency (for review, see Refs. 45 and 46).

In the present study, we used direct plasmid injection to examine whether the utrophin 3'-UTR contains elements that may confer differential stability to reporter transcripts. Our data show that the utrophin 3'-UTR can indeed increase the levels of chimeric LacZ mRNAs in soleus muscles by an extent similar to that seen for endogenous utrophin transcripts. This suggests, therefore, that although the 5'-UTR and the coding regions can also contribute to the stability of specific mRNAs (45, 46), it appears that in the case of utrophin transcripts the 3'-UTR is primarily responsible for controlling their turnover rate. This view is, in fact, further supported by the observation that expression of the A and B transcripts, which differ mostly in their 5'-UTR (8), are both increased in soleus muscles.

In an attempt to determine whether the difference in utrophin mRNA stability in fast vs. slow muscles involves differential expression of RNA-binding proteins, we performed UV-crosslinking experiments using cytoplasmic extracts from soleus and EDL muscles. Our results indicate that both muscle types express a similar pattern of proteins that can bind to the utrophin 3'-UTR. However, we noted in these experiments a clear difference in the relative content of these RNA-binding proteins. Indeed, the extract from fast EDL muscles appears to contain significantly more protein factors that interact with the utrophin 3'-UTR. This would indicate that these complexes represent destabilizing factors because EDL muscle expresses fewer utrophin transcripts. In this context, it is interesting to note that the utrophin 3'-UTR contains six AUREs (Gramolini, Bélanger, and Jasmin, unpublished observations). It is therefore possible that these elements may preferentially destabilize utrophin transcripts in fast muscle via interactions with members of the AURE-binding protein family. Experiments are currently under way to see whether expression of these RNA-binding proteins differ between fast and slow muscles.

A key issue that also deserves further attention deals with the nature of the signaling events that ultimately control expression of these RNA-binding proteins. In this context, we have recently shown that elimination of electrical activity via surgical denervation induces an increase in the amount of cytoplasmic factors that interact with the 3'-UTR of AChE (5). In addition, Booth and colleagues (59) have previously demonstrated that increased contractile activity induces a significant decrease in RNA-protein interactions in the 3'-UTR of cytochrome c mRNA. On the basis of these observations, it appears that electrical activity represents an initiating signal that ultimately influences, via yet to be determined signaling cascades, the abundance of RNA-binding proteins, which, in turn, affect the stability of specific transcripts. Accordingly, it is tempting to speculate that the differences in the amount and pattern of electrical activity, known to exist between soleus and EDL muscles (27), are responsible for controlling the levels of cytoplasmic factors that interact with the utrophin 3'-UTR. In this scenario, calcium may constitute a key signal, linking membrane events to changes in mRNA expression, because the continuous activation pattern of soleus muscles results in sustained elevations of the intracellular concentration of calcium.

Recent studies have led to the idea that calcineurin, a calcium-regulated protein phosphatase, may serve as a key player in the control of the slow muscle phenotype by acting via nuclear factor of activated T cells (NFAT) transcription factors (for review, see Ref. 43). Interestingly, calcineurin has also been implicated in the regulation of mRNA stability in different cell types. In skeletal muscle for example, Taylor and colleagues (36) have shown that calcineurin can modulate the stability of AChE transcripts in differentiating C2C12 cells maintained in culture. Along those lines, we have recently observed that overexpression of a constitutively active form of calcineurin in transgenic mice results in an increased expression of utrophin in soleus muscles (13). Together with the findings of the present study, these data suggest that the greater stability of utrophin transcripts in soleus muscles may result from the sustained intracellular levels of calcium seen in soleus muscles, which, in turn, activate calcineurin, thereby regulating the stability of utrophin mRNAs.

The results of the present study showing that slow muscles contain more utrophin may explain, in fact, why fast fibers are preferentially affected in DMD patients (56). Indeed, in the absence of dystrophin, extrasynaptic utrophin may help protect slow fibers against the damage elicited by repetitive mechanical stress. In addition, our findings that utrophin mRNAs appear more stable in slow muscles provide an in vivo model system with which to begin characterizing precisely cis-acting elements and trans-activating factors that control the half-life of utrophin transcripts in skeletal muscle fibers. The results of the present studies should therefore lead to the identification of novel posttranscriptional targets for which pharmacological manipulations may be envisaged to ultimately increase the endogenous levels of utrophin in DMD muscle fibers. Accordingly, experiments remain to be done that will nicely complement ongoing studies aimed at increasing utrophin levels via interventions on transcriptional events.


    ACKNOWLEDGEMENTS

We thank J. A. Lunde for expert technical assistance and D. J. Parry for helpful discussion.


    FOOTNOTES

This work was supported by grants from the Association Française Contre les Myopathies, the Muscular Dystrophy Association of America, and the Canadian Institutes of Health Research (CIHR). During the course of this work, A. O. Gramolini was supported by a Strategic Area of Development Fellowship from the University of Ottawa and is now supported by a Postdoctoral Fellowship from CIHR. B. J. Jasmin is a CIHR Investigator.

Present address of A. O. Gramolini: Howard Hughes Medical Institute and Dept. of Cell Biology, Duke Univ. Medical Center, Durham, NC 27710.

Address for reprint requests and other correspondence: B. J. Jasmin, Dept. of Cellular and Molecular Medicine, Faculty of Medicine, Univ. of Ottawa, 451 Smyth Rd., Ottawa, Ontario, Canada K1H 8M5 (E-mail: jasmin{at}uottawa.ca).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 15 March 2001; accepted in final form 5 June 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Angus, LM, Chan RYY, and Jasmin BJ. Role of intronic N- and E-boxes in the transient transcriptional induction of the acetylcholinesterase gene during myogenic differentiation. J Biol Chem 276: 17603-17609, 2001[Abstract/Free Full Text].

2.   Bag, J, and Wu J. Translational control of poly(A)-binding protein expression. Eur J Biochem 237: 143-152, 1996[Abstract].

3.   Bélanger, G, Chan RYY, and Jasmin BJ. Cis- and trans-acting factors controlling acetylcholinesterase mRNA stability during myogenic differentiation. Soc Neurosci Abstr 1089: 26, 2000.

4.   Blake, DJ, Tinsley JM, and Davies KE. Utrophin: a structural and functional comparison to dystrophin. Brain Pathol 6: 37-47, 1996[ISI][Medline].

5.   Boudreau-Lariviere, C, Chan RY, Wu J, and Jasmin BJ. Molecular mechanisms underlying the activity-linked alterations in acetylcholinesterase mRNAs in developing versus adult rat skeletal muscles. J Neurochem 74: 2250-2258, 2000[ISI][Medline].

6.   Boudreau-Lariviere, C, Parry DJ, and Jasmin BJ. Myotubes originating from single fast and slow satellite cells display similar patterns of AChE expression. Am J Physiol Regulatory Integrative Comp Physiol 278: R140-R148, 2000[Abstract/Free Full Text].

7.   Burden, SJ. The formation of neuromuscular synapses. Genes Dev 12: 133-148, 1998[Free Full Text].

8.   Burton, EA, Tinsley JM, Holzfeind PJ, Rodrigues NR, and Davies KE. A second promoter provides an alternative target for therapeutic upregulation of utrophin in Duchenne muscular dystrophy. Proc Natl Acad Sci USA 96: 14025-14030, 1999[Abstract/Free Full Text].

9.   Chan, RY, Adatia FA, Krupa AM, and Jasmin BJ. Increased expression of acetylcholinesterase T and R transcripts during hematopoietic differentiation is accompanied by parallel elevations in the levels of their respective molecular forms. J Biol Chem 273: 9727-9733, 1998[Abstract/Free Full Text].

10.   Chan, RY, Boudreau-Lariviere C, Angus LM, Mankal FA, and Jasmin BJ. An intronic enhancer containing an N-box motif is required for synapse- and tissue-specific expression of the acetylcholinesterase gene in skeletal muscle fibers. Proc Natl Acad Sci USA 96: 4627-4632, 1999[Abstract/Free Full Text].

11.   Chopard, A, Pons F, Charpiot P, and Marini JF. Quantitative analysis of relative protein contents by Western blotting: comparison of three members of the dystrophin-glycoprotein complex in slow and fast rat skeletal muscle. Electrophoresis 21: 517-522, 2000[ISI][Medline].

12.   Day, DA, and Tuite MF. Post-transcriptional gene regulatory mechanisms in eukaryotes: an overview. J Endocrinol 157: 361-371, 1998[Abstract/Free Full Text].

13.   Deschenes, JL, Viau F, Chin E, Michel RN, and Jasmin BJ. Role of calcineurin in the regulation of synaptic proteins in skeletal muscle. Soc Neurosci Abstr 26: 1090, 2000.

14.   Duclert, A, and Changeux JP. Acetylcholine receptor gene expression at the developing neuromuscular junction. Physiol Rev 75: 339-368, 1995[Free Full Text].

15.   Emery, AE. Population frequencies of inherited neuromuscular diseases---a world survey. Neuromuscul Disord 1: 19-29, 1991[Medline].

16.   Forster, E, Otten U, and Frotscher M. Developmental neurotrophin expression in slice cultures of rat hippocampus. Neurosci Lett 155: 216-219, 1993[ISI][Medline].

17.   Fuentes, ME, and Taylor P. Control of acetylcholinesterase gene expression during myogenesis. Neuron 10: 679-687, 1993[ISI][Medline].

18.   Gramolini, AO, Angus LM, Schaeffer L, Burton EA, Tinsley JM, Davies KE, Changeux JP, and Jasmin BJ. Induction of utrophin gene expression by heregulin in skeletal muscle cells: role of the N-box motif and GA binding protein. Proc Natl Acad Sci USA 96: 3223-3227, 1999[Abstract/Free Full Text].

19.  Gramolini AO, Bélanger G, and Jasmin BJ. Distinct regions in the 3'UTR are responsible for targeting and stabilizing utrophin transcripts in skeletal muscle cells. J Cell Biol. In press.

20.   Gramolini, AO, Burton EA, Tinsley JM, Ferns MJ, Cartaud A, Cartaud J, Davies KE, Lunde JA, and Jasmin BJ. Muscle and neural isoforms of agrin increase utrophin expression in cultured myotubes via a transcriptional regulatory mechanism. J Biol Chem 273: 736-743, 1998[Abstract/Free Full Text].

21.   Gramolini, AO, Dennis CL, Tinsley JM, Robertson GS, Cartaud J, Davies KE, and Jasmin BJ. Local transcriptional control of utrophin expression at the neuromuscular synapse. J Biol Chem 272: 8117-8120, 1997[Abstract/Free Full Text].

22.   Gramolini, AO, and Jasmin BJ. Expression of the utrophin gene during myogenic differentiation. Nucleic Acids Res 27: 3603-3609, 1999[Abstract/Free Full Text].

23.   Gramolini, AO, Karpati G, and Jasmin BJ. Discordant expression of utrophin and its transcript in human and mouse skeletal muscles. J Neuropathol Exp Neurol 58: 235-244, 1999[ISI][Medline].

24.   Gramolini, AO, Wu J, and Jasmin BJ. Regulation and functional significance of utrophin expression at the mammalian neuromuscular synapse. Microsc Res Tech 49: 90-100, 2000[ISI][Medline].

25.   Guo, WX, Nichol M, and Merlie JP. Cloning and expression of full length mouse utrophin: the differential association of utrophin and dystrophin with AChR clusters. FEBS Lett 398: 259-264, 1996[ISI][Medline].

26.   Helliwell, TR, Man NT, Morris GE, and Davies KE. The dystrophin-related protein, utrophin, is expressed on the sarcolemma of regenerating human skeletal muscle fibres in dystrophies and inflammatory myopathies. Neuromuscul Disord 2: 177-184, 1992[Medline].

27.   Hennig, R, and Lomo T. Firing patterns of motor units in normal rats. Nature 314: 164-166, 1985[ISI][Medline].

28.   Ho-Kim, MA, and Rogers PA. Quantitative analysis of dystrophin in fast- and slow-twitch mammalian skeletal muscle. FEBS Lett 304: 187-191, 1992[ISI][Medline].

29.   Jasmin, BJ, Alameddine H, Lunde JA, Stetzkowski-Marden F, Collin H, Tinsley JM, Davies KE, Tome FM, Parry DJ, and Cartaud J. Expression of utrophin and its mRNA in denervated mdx mouse muscle. FEBS Lett 374: 393-398, 1995[ISI][Medline].

30.   Karpati, G, Carpenter S, Morris GE, Davies KE, Guerin C, and Holland P. Localization and quantitation of the chromosome 6-encoded dystrophin-related protein in normal and pathological human muscle. J Neuropathol Exp Neurol 52: 119-128, 1993[ISI][Medline].

31.   Khurana, TS, Rosmarin AG, Shang J, Krag TO, Das S, and Gammeltoft S. Activation of utrophin promoter by heregulin via the ets-related transcription factor complex GA-binding protein alpha/beta. Mol Biol Cell 10: 2075-2086, 1999[Abstract/Free Full Text].

32.   Khurana, TS, Watkins SC, Chafey P, Chelly J, Tome FM, Fardeau M, Kaplan JC, and Kunkel LM. Immunolocalization and developmental expression of dystrophin related protein in skeletal muscle. Neuromuscul Disord 1: 185-194, 1991[Medline].

33.   Krejci, E, Legay C, Thomine S, Sketelj J, and Massoulie J. Differences in expression of acetylcholinesterase and collagen Q control the distribution and oligomerization of the collagen-tailed forms in fast and slow muscles. J Neurosci 19: 10672-10679, 1999[Abstract/Free Full Text].

34.   Kues, WA, Sakmann B, and Witzemann V. Differential expression patterns of five acetylcholine receptor subunit genes in rat muscle during development. Eur J Neurosci 7: 1376-1385, 1995[ISI][Medline].

35.   Liebhaber, SA. mRNA stability and the control of gene expression. Nucleic Acids Symp Ser 36: 29-32, 1997[Medline].

36.   Luo, ZD, Wang Y, Werlen G, Camp S, Chien KR, and Taylor P. Calcineurin enhances acetylcholinesterase mRNA stability during C2-C12 muscle cell differentiation. Mol Pharmacol 56: 886-894, 1999[Abstract/Free Full Text].

37.   Matsumura, K, and Campbell KP. Dystrophin-glycoprotein complex: its role in the molecular pathogenesis of muscular dystrophies. Muscle Nerve 17: 2-15, 1994[ISI][Medline].

38.   Menold, MM, and Repasky EA. Heterogeneity of spectrin distribution among avian muscle fiber types. Muscle Nerve 7: 408-414, 1984[ISI][Medline].

39.   Michel, RN, Vu CQ, Tetzlaff W, and Jasmin BJ. Neural regulation of acetylcholinesterase mRNAs at mammalian neuromuscular synapses. J Cell Biol 127: 1061-1069, 1994[Abstract].

40.   Mizuno, Y, Yoshida M, Nonaka I, Hirai S, and Ozawa E. Expression of utrophin (dystrophin-related protein) and dystrophin-associated glycoproteins in muscles from patients with Duchenne muscular dystrophy. Muscle Nerve 17: 206-216, 1994[ISI][Medline].

41.   Newey, SE, Gramolini AO, Wu J, Holzfeind P, Jasmin BJ, Davies KE, and Blake DJ. A novel mechanism for modulating synaptic gene expression: Differential localization of alpha -dystrobrevin transcripts in skeletal muscle. Mol Cell Neurosci 17: 127-140, 2001[ISI][Medline].

42.   Nguyen, TM, Ellis JM, Love DR, Davies KE, Gatter KC, Dickson G, and Morris GE. Localization of the DMDL gene-encoded dystrophin-related protein using a panel of nineteen monoclonal antibodies: presence at neuromuscular junctions, in the sarcolemma of dystrophic skeletal muscle, in vascular and other smooth muscles, and in proliferating brain cell lines. J Cell Biol 115: 1695-1700, 1991[Abstract].

43.   Olson, EN, and Williams RS. Remodeling muscles with calcineurin. Bioessays 22: 510-519, 2000[ISI][Medline].

44.   Pette, D, and Staron RS. Mammalian skeletal muscle fiber type transitions. Int Rev Cytol 170: 143-223, 1997[Medline].

45.   Rajagopalan, LE, and Malter JS. Regulation of eukaryotic messenger RNA turnover. Prog Nucleic Acid Res Mol Biol 56: 257-286, 1997[ISI][Medline].

46.   Ross, J. Control of messenger RNA stability in higher eukaryotes. Trends Genet 12: 171-175, 1996[ISI][Medline].

47.   Sanes, JR, and Lichtman JW. Development of the vertebrate neuromuscular junction. Annu Rev Neurosci 22: 389-442, 1999[ISI][Medline].

48.   Schiaffino, S, and Reggiani C. Molecular diversity of myofibrillar proteins: gene regulation and functional significance. Physiol Rev 76: 371-423, 1996[Abstract/Free Full Text].

49.   Sigrist, SJ, Thiel PR, Reiff DF, Lachance PE, Lasko P, and Schuster CM. Postsynaptic translation affects the efficacy and morphology of neuromuscular junctions. Nature 405: 1062-1065, 2000[ISI][Medline].

50.   Takemitsu, M, Ishiura S, Koga R, Kamakura K, Arahata K, Nonaka I, and Sugita H. Dystrophin-related protein in the fetal and denervated skeletal muscles of normal and mdx mice. Biochem Biophys Res Commun 180: 1179-1186, 1991[ISI][Medline].

51.   Talmadge, RJ, Roy RR, and Edgerton VR. Muscle fiber types and function. Curr Opin Rheumatol 5: 695-705, 1993[Medline].

52.   Taormino, JP, and Fambrough DM. Pre-translational regulation of the (Na+-K+)-ATPase in response to demand for ion transport in cultured chicken skeletal muscle. J Biol Chem 265: 4116-4123, 1990[Abstract/Free Full Text].

53.   Tinsley, JM, Blake DJ, Roche A, Fairbrother U, Riss J, Byth BC, Knight AE, Kendrick-Jones J, Suthers GK, Love DR, Edwards YH, and Davies KE. Primary structure of dystrophin-related protein. Nature 360: 591-593, 1992[ISI][Medline].

54.   Tinsley, JM, Potter AC, Phelps SR, Fisher R, Trickett JI, and Davies KE. Amelioration of the dystrophic phenotype of mdx mice using a truncated utrophin transgene. Nature 384: 349-353, 1996[ISI][Medline].

55.   Vater, R, Young C, Anderson LV, Lindsay S, Blake DJ, Davies KE, Zuellig R, and Slater CR. Utrophin mRNA expression in muscle is not restricted to the neuromuscular junction. Mol Cell Neurosci 10: 229-242, 1998[ISI].

56.   Webster, C, Silberstein L, Hays AP, and Blau HM. Fast muscle fibers are preferentially affected in Duchenne muscular dystrophy. Cell 52: 503-513, 1988[ISI][Medline].

57.   Williams, MW, Resneck WG, and Bloch RJ. Membrane skeleton of innervated and denervated fast- and slow-twitch muscle. Muscle Nerve 23: 590-599, 2000[ISI][Medline].

58.   Worton, R. Muscular dystrophies: diseases of the dystrophin-glycoprotein complex. Science 270: 755-756, 1995[Free Full Text].

59.   Yan, Z, Salmons S, Dang YI, Hamilton MT, and Booth FW. Increased contractile activity decreases RNA-protein interaction in the 3'-UTR of cytochrome c mRNA. Am J Physiol Cell Physiol 271: C1157-C1166, 1996[Abstract/Free Full Text].


Am J Physiol Cell Physiol 281(4):C1300-C1309
0363-6143/01 $5.00 Copyright © 2001 the American Physiological Society