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Human Molecular Genetics, 2003, Vol. 12, No. 10 1087-1099
DOI: 10.1093/hmg/ddg133
© 2003 Oxford University Press

Restoration of dystrophin expression in mdx muscle cells by chimeraplast-mediated exon skipping

Carmen Bertoni1, Catherine Lau1 and Thomas A. Rando1,2,*

1Department of Neurology and Neurological Sciences, Stanford University School of Medicine, Stanford, CA, USA and 2GRECC, VA Palo Alto Health Care System, Palo Alto, CA, USA

Received December 15, 2002; Accepted March 18, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The most common types of dystrophin gene mutations that cause Duchenne muscular dystrophy (DMD) are large deletions that result in a shift of the translational reading frame. Such mutations generally lead to a complete absence of dystrophin protein in the muscle cells of affected individuals. Any therapeutic modality that could restore the reading frame would have the potential to substantially reduce the severity of the disease by allowing the production of an internally deleted, but partially functional, dystrophin protein as occurs in Becker muscular dystrophy (BMD). One approach to restoring the reading frame would be to alter the splicing of the pre-mRNA to produce an in-frame transcript. We have tested the ability of chimeric RNA/DNA oligonucleotides (chimeraplasts) to alter key bases in specific splice sequences in the dystrophin gene to induce exon skipping. Using cells from the mdx mouse as a model system, we show that chimeraplast-mediated base conversion in the intron 22/exon 23 splice junction induces alternative splicing and the production of in-frame transcripts. Interestingly, multiple alternative transcripts were induced by this targeted splice site mutation. Direct sequencing indicated that several of these were predicted to produce in-frame dystrophin transcripts with internal deletions. Indeed, multiple forms of dystrophin protein were observed by western blot analysis, and the functionality of the products was demonstrated by the restoration of expression and localization of a dystrophin-associated protein, {alpha}-dystroglycan, in differentiated cells. These data demonstrate that chimeraplasts can induce exon skipping by altering splice site sequences at the genomic level. As such, chimeraplast-mediated exon skipping has the potential to be used to transform a severe DMD phenotype into a much milder BMD phenotype.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Duchenne muscular dystrophy (DMD) and Becker muscular dystrophy (BMD) are X-linked muscle diseases caused by defects in the 2.5 Mb gene that encodes the dystrophin protein (1). More than 80% of patients with DMD or BMD have large deletions in the dystrophin gene, with duplications, small deletions, insertions and point mutations making up the remainder of the cases (2). In general, the severe DMD phenotype is caused by mutations, such as frame-shift deletions and nonsense point mutations, that lead to complete dystrophin deficiency. The milder BMD phenotype is usually correlated with deletions that preserve the translational reading frame and produce internally deleted, but partially functional, dystrophin proteins (3). Thus, any therapeutic modality that could convert a frame-shift mutation to a non-frame-shift mutation has the potential to convert a severe DMD phenotype into a milder BMD phenotype.

The most widely studied strategies to restore the expression of dystrophin in dystrophin-deficient muscles involve the use of viral-mediated delivery to introduce a functional dystrophin cDNA (4–7). This approach has been effective in animal models and holds promise for application in humans. Furthermore, this approach is applicable to the treatment of any type of DMD mutation, not just frame-shift deletions. However, viral delivery systems face challenges inherent to immune responses against the vectors, issues related to viral integration into the genome, and the appropriate regulation of expression of the delivered gene. Several groups have investigated the possibility of using antisense oligonucleotides to alter the processing of the dystrophin transcript to induce exon skipping and convert frame-shift mutations into non-frame-shift mutations (8–12). This technology avoids some of the hurdles of viral mediated gene therapy. However, this approach would require repeated delivery of oligonucleotides in order to maintain dystrophin expression, or there would be a need for sustained expression of an antisense transcript as could be achieved with a viral delivery system (8). An appealing alternative to antisense-mediated exon skipping to restore the translational reading frame would be a gene modification strategy that would be permanent. Current gene repair strategies that could be applied to this problem include technologies involving chimeric oligonucleotides, single-stranded DNA oligonucleotides, and short-fragment homologous recombination (13,14).

Gene repair involving chimeric RNA/DNA oligonucleotides (chimeraplasts) is based upon the activity of these vectors to induce base pair conversions at the genomic level (15). Their mechanism of action involves the utilization of innate mismatch repair activities in the cell, directing the substitution of a single base in a specific DNA sequence (16,17). Chimeraplast-mediated gene repair has been successfully applied to different cell types, both in vitro and in vivo (18–25). Although primarily studied as an application for repairing point mutations, chimeraplast-mediated base conversions of consensus splice site sequences also have the potential to alter splicing and thus restore the translational reading frame in a gene with a frame-shift mutation.

In this report, we have investigated the possibility of using chimeraplasts to alter the splicing of the dystrophin gene. The model system, the mdx mouse, has a nonsense mutation in exon 23 of the dystrophin gene that leads to complete absence of the dystrophin protein. We have previously shown that chimeraplasts are able to correct this point mutation both in vitro and in vivo (20,21). We have tested whether targeted mutation of a single base in the intron 22/exon 23 splice junction could alter the splicing of the dystrophin transcript to skip exon 23. The predicted product in which exon 22 is directly spliced to exon 24 would be in-frame and produce a nearly full-length dystrophin protein. Although the mdx mouse does not have frame-shift deletion, it does provides an opportunity to investigate how stable, single-base alterations in genomic DNA may be used to redirect splicing that would be applicable to frame-shift mutations in the dystrophin gene. We have found that altering the intron 22/exon 23 splice junction does yield in-frame transcripts and shortened forms of the dystrophin protein. These studies expand the therapeutic application of chimeraplast-mediated single base substitutions for frame-shift deletions.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
A targeting chimeraplast, termed MDX3 (Fig. 1A), was designed to induce a G-to-T conversion in the exon 23 splice acceptor site of the intron 22/exon 23 splice junction of the mouse dystrophin gene (Fig. 1B). Alteration of the base targeted by MDX3 would result in the disruption of the intron 22/exon 23 consensus sequence and create a novel restriction site in this position (Fig. 1B). The predicted sequence of a transcript resulting from the splicing of exon 22 to exon 24 would be in-frame and would be translated into a protein 71 amino acids shorter than full-length dystrophin. As a control, we used a chimeric oligonucleotide, MDX4 (Fig. 1A), designed to be perfectly complementary to the intron 22/exon 23 consensus sequence and thus to have no activity to induce alterations in the genomic sequence.



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  		Figure 1. Chimeraplast design and targeting strategy. (A) Linear sequences of targeting (MDX3) and control (MDX4) chimeraplasts are shown, with the secondary hairpin-loop structures shown below. Upper case letters indicate DNA residues and lower case letters indicate 2'-O-methyl RNA residues. The single base pair difference between MDX3 and MDX4 is indicated in the shaded box. Underlined sequences indicate the region of homology with the intron 22/exon 23 splice site of the dystrophin gene. BLAST search of the mouse genome database confirmed the specificity for the region of the dystrophin sequence targeted by the chimeraplasts. (B) The sequence of the dystrophin gene flanking the intron 22/exon 23 splice junction is shown. The mdx mutation is located 48 bp downstream of the splice site at position 3185. The nucleotides underlined correspond to the sequence used to design the targeting and control chimeraplasts. MDX3 is designed to induce a G-to-T conversion in the acceptor splice site of exon 23 at base 3137 of the dystrophin gene. The single base pair substitution would create a novel Hinf I restriction site (gray box).

 
Evidence of chimeraplast-mediated single base substitution at the genomic level
The specificity of MDX3 to induce the G-to-T conversion in the intron 22/exon 23 splice site was assessed using an ARMS assay of genomic DNA. Cells were maintained in culture for at least 15 days after chimeraplast transfection. Using PCR primers designed to amplify the sequence bearing the G-to-T transversion, we observed a specific amplification product only in cells transfected with the targeting chimeraplast (Fig. 2A). Cells transfected with control chimeraplast or untransfected cells failed to produce any amplification product using these primers.



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  		Figure 2. Evidence of chimeraplast mediated gene conversion. (A) Genomic DNA samples isolated from untreated mdx cells or mdx cells transfected with control (MDX4) or targeting (MDX3) chimeraplasts were amplified using primers flanking the intron 22/exon 23 splice site sequence. PCR reactions using the ‘w’ primer (see Methods) show a specific 100 bp product corresponding to the intron 22/exon 23 splice site sequence in all samples. PCR reactions using the ‘m3’ primer were carried out to identify intron 22/exon 23 splice site sequences containing the G-to-T conversion shown in Figure 1B. Amplification with this primer was seen only in cells transfected with MDX3. Molecular weight markers are shown in the lane labeled ‘M’, and the last two lanes on the right side of the gel are primer controls lacking genomic template. (B) Genomic DNA was extracted after chimeraplast transfection and amplified using primers located 50 bp upstream or 100 bp downstream the intron 22/exon 23 splice junction. The PCR product was digested with Hinf I to confirm the presence of the single base mutation created by MDX3 (U: undigested product). (C) The G-to-T transversion at the intron 22/exon 23 splice junction was detected by direct sequencing of PCR amplified products of DNA isolated from MDX3-transfected cells. The single base pair substitution was detectable after the first round of amplification as a minor component, consistent with the conversion occurring in only a subset of cells. The second round of amplification clearly demonstrates the specific targeted mutation by MDX3.

 
The G-to-T conversion by the targeting chimeraplast was also confirmed by direct digestion of PCR amplification products (Fig. 2B). Transfected cells were maintained in culture for more than 45 days and genomic DNA was extracted. Specific PCR primers were used to produce an amplification product of 150 bp that contained the intron 22/exon 23 splice junction which was confirmed by direct sequencing. After purification, the PCR products were subjected to digestion using Hinf I restriction endonuclease. Digestion products at the expected size of ~50 and 100 bp were observed from cells transfected with MDX3 (Fig. 2B), demonstrating that the targeting chimeraplast had induced the single base pair substitution.

To analyze further for evidence of gene modification at the genomic level, we directly sequenced PCR-amplification products of the intron 22/exon 23 splice junction from MDX3-transfected and control cells. The G-to-T conversion could be detected after one round of PCR amplification, and confirmed after a second round of amplification, only in MDX3-transfected cells (Fig. 2C). These results were reproduced in 10 independent experiments and were confirmed by sequence analysis using both forward and reverse primers. These results confirm the specific base alteration by the targeting chimeraplast in a subset of the transfected cells at the genomic level.

Dystrophin protein expression in chimeraplast-treated mdx muscle cells
The restoration of dystrophin protein expression was assessed 3 weeks after transfection and 24–96 h after induction of differentiation as previously described (20). Western blotting using an antibody to the rod domain of the dystrophin protein demonstrated at least two distinct protein products whose molecular weights were less than that of full-length dystrophin (Fig. 3A). An antibody directed toward the C-terminal domain confirmed the expression of these two products but also revealed the presence of a third protein product of even lower molecular weight (Fig. 3A). Based on the antibody specificity, the larger products contained the exon 31/32 region of the protein whereas the smaller product did not. It is possible to obtain only a rough approximation of molecular weights of these products given their size and limited mobility. All proteins ran faster than dystrophin or utrophin, thus establishing 400 kDa as the upper limit of their sizes, and all proteins ran slower than the highest molecular weight marker of 200 kDa, thus establishing a lower limit. The two higher molecular weight bands (Fig. 3A) ran closer to the 400 kDa end of the spectrum and the lower molecular weight band (recognized only by the antibody against the C-terminal region of the protein) ran closer to the 200 kDa end. This pattern of dystrophin proteins was obtained in five independent experiments and was confirmed by immunoprecipitation analysis. Furthermore, this pattern of expression was present in myotube cultures derived from MDX3-transfected myoblasts that had been maintained in culture for up to 3 months after transfection, confirming that the effect of the chimeraplast treatment was stably inherited.



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  		Figure 3. Analysis of dystrophin expression in chimeraplast-treated cells. (A) Myoblast cultures transfected with MDX3 or MDX4 chimeraplasts were induced to differentiate and then analyzed for the expression of dystrophin. Western blot analysis using an antibody (Mandys-8) directed toward the rod domain of the dystrophin protein revealed the presence of at least two different protein products of slightly different molecular weights and both smaller than full-length dystrophin. The same blot re-probed using an antibody (Dys-2) specific for the C-terminal domain of dystrophin showed the presence of a third band of lower molecular weight. Untreated cells or cells treated with control chimeraplast showed no detectable expression of dystrophin protein. The bottom of the gel in these panels corresponds to a molecular weight marker of ~200 kDa. (B) Proliferating mdx myoblasts transfected with the targeting chimeraplast were maintained in culture for 2 months after transfection prior to the induction of differentiation. Expression of dystrophin is evident in clusters of myotubes 48 h after the induction of differentiation using an antibody directed against the C-terminal domain of dystrophin (Dys-2). Two different clusters of dystrophin-expressing myotubes are shown. (C) Dystrophin immunostaining is shown using antibodies directed toward three different regions of the dystrophin protein: the exon 26 region (Mandys-18), the exon 31/32 region (Mandys-8), and the C-terminal region (Dys-2). Clusters of dystrophin-expressing myotubes were easily identified with the Mandys-8 and Dys-2 antibodies, but no immunoreactivity was seen in any myotubes in the cultures when the antibody against exon 26 (Mandys-18) was used, suggesting that all forms of dystrophin produced skipped this exon. No dystrophin immunoreactivity was seen in MDX4-transfected cells.

 
We have previously used western blot analysis to quantitate the level of dystrophin expression in cultures of mdx cells transfected with chimeraplasts, thus providing a measure of efficacy of gene conversion (20). In this current study, however, the presence of multiple bands on the western blots made this measure of efficacy more difficult. As an alternative approach, we constructed a standard curve using a mixture of extracts from C57 and mdx myotubes to determine the limit of our ability to detect dystrophin protein expression by western blot analysis. According to these studies, dystrophin could be detected as long as the percentage of dystrophin-expressing cells was greater than 1%. Thus we concluded that the efficacy of single base substitution mediated by the MDX3 chimeraplast was in the range of 1–5%. This range is similar to that previously reported for a chimeraplast designed to correct the mdx mutation (20,21) and is in the range of chimeraplast-mediated base conversion shown by others (19,23,26).

The expression of some form of the dystrophin protein in a subset of MDX3-transfected cells was also confirmed by immunostaining myotube cultures. Consistent with the western blot results, we observed clusters of dystrophin-positive fibers after differentiation only in MDX3-transfected cells (Fig. 3B). Immunostaining of treated cells with exon-specific anti-dystrophin antibodies confirmed the presence of internally deleted forms of dystrophin in myotubes derived from MDX3-transfected cells (Fig. 3C), whereas myotube cultures derived from untransfected cells or from MDX4-transfected cells did not stain positively with any of the antibodies. In myotube cultures derived from MDX3-transfected cells, clusters of myotubes stained positively with antibodies against the C-terminal domain and the exon 31/32 region of the dystrophin protein, but none of the myotubes stained positively with an antibody against exon 26 (Fig. 3C). These data are consistent with the western blot data, indicating that alteration of the intron 22/exon 23 splice site results in the skipping of multiple exons, not just exon 23 (Fig. 3A). Multiple exon skipping was also observed by other investigators using antisense oligonucleotides to interfere with the intron 22/exon 23 splice junction (9).

Restoration of the {alpha}-dystroglycan expression
It has been shown that the expression of {alpha}-dystroglycan ({alpha}-DG), a major component of the dystrophin–glycoprotein complex, is reduced by 80–90% in the mdx mouse due to the lack of dystrophin (27). We therefore used immunostaining to determine if the dystrophin proteins produced by the disruption of the intron 22/exon 23 splice junction could restore the expression of {alpha}-DG. Immunostaining of MDX3-transfected cultures that had been differentiated for 4 days confirmed the localization of {alpha}-DG at the sarcolemma membrane in clusters of myotubes (Fig. 4). These results clearly demonstrate that at least one of the dystrophin proteins produced was functional, consistent with the functional properties of engineered dystrophin proteins lacking large portions of the rod domain (28).



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  		Figure 4. Restoration of the expression of {alpha}-dystroglycan after MDX3 treatment. Myotube cultures were generated from myoblasts that had been transfected with MDX3 or MDX4 chimeraplasts 4 weeks earlier. The cultures were analyzed for the expression of {alpha}-DG by immunofluorescence. Four days after the induction of differentiation, myotubes derived from cells transfected with MDX3 showed membrane localization of {alpha}-DG, whereas no {alpha}-DG could be detected in myotubes derived from MDX4-transfected cells.

 
Detection of the alternative spliced products induced by MDX3
Based on the immunostaining and the immunoblot analysis using exon-specific antibodies, it was clear that we were identifying products of alternate splicing other than just that due to the skipping of exon 23. We used RT–PCR analysis to identify alternatively spliced products induced by the disruption of the intron 22/exon 23 splice site in cells that had been maintained in culture 2 weeks after chimeraplast transfection. We designed specific primers spanning exon 22 to exon 32 of the dystrophin gene. Products representing the full-length transcripts were amplified from untreated and chimeraplast-treated cells using these primers (Fig. 5). We were unable to detect any alternatively spliced transcripts within this region in cells transfected with the targeting chimeraplast (Fig. 5). The absence of alternative splicing products was also confirmed by the introduction of an additional step of nested RT–PCR amplification using inner primers (data not shown).



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  		Figure 5. Analysis of the dystrophin transcripts in chimeraplast-transfected cells. Cells transfected with MDX3 chimeraplast were induced to differentiate two weeks after transfection and analyzed for the expression of alternatively spliced transcripts. Total messenger RNA was retrotranscribed and PCR amplified using a reverse primer in exon 33 (R33) and a forward primer in exon 22 (F22) of the dystrophin gene. Nested PCR was conducted using a forward primer located in exon 22 (F22in) and reverse primers in exons 23 (R23), 24 (R24), 25 (R25), 30 (R30) and 32 (R32), as indicated. Amplification within this region failed to reveal any alternative spliced dystrophin transcript resulting from the skipping of exons between 23 and 32 in MDX3-transfected cells compared with untransfected cells. PCR product sizes are indicated on the left side of the gel. Molecular weight markers are shown in the lanes labeled ‘M’.

 
We therefore extended our investigation to a larger portion of the dystrophin gene using primers that spanned from exon 9 to exon 50 (Fig. 6A), thus covering almost 50% of the entire dystrophin cDNA. A second PCR amplification was performed using primer pairs (see Materials and Methods and Fig. 6A) located within the region of the dystrophin transcript amplified in the first step. We observed five products (labeled ‘b’, ‘c’, ‘d’, ‘e’ and ‘f’) in MDX3-transfected cells (Fig. 6B and C), and this pattern was also observed in cells that had been maintained in culture for 3 months after transfection. None of these products were ever detected in untransfected cells or cells transfected with the control chimeraplast. Direct sequencing of these products showed that they represented five different splice variants, each skipping exon 23 plus additional exons both upstream and downstream of the intron 22/exon 23 splice junction, with as many as 26 exons (exons 12–37) being skipped in the smallest product (Fig. 6C and D). Three out of the five splice variants (products ‘c’, ‘d’ and ‘f’) were in-frame (Table 1). The predicted amino acid sequences from these in-frame transcripts would comprise proteins between 271 and 350 kDa (Table 1), consistent with the range suggested by western blot analysis (Fig. 3A).



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  		Figure 6. Alternative splicing induced by MDX3 chimeraplast. (A) Schematic representation of the strategy used for the detection of alternatively spliced products. Nested RT–PCR reactions were performed between exon 8 and exon 50 of the dystrophin cDNA. First-strand retrotranscriptions and amplifications were performed using a forward primer in exon 8 (F8) paired with reverse primers in exons 33 (R33), 35 (R35out), 43 (R43out), and 50 (R50out). The size of each first amplification product is shown to the right. Nested PCR was then conducted using forward primers in exons 9 (F9) or 19 (F19) paired with reverse primers in exons 32 (R32), 35 (R35in), 43 (R43in), and 50 (R50in). (B) Cells transfected with MDX3 or MDX4 chimeraplasts were analyzed for alternative splicing 15 days after transfection and 3 days after the induction of differentiation. Alternative splicing was analyzed by nested RT–PCR using a forward primer in exon 19 (F19) and a reverse primer in exon 32 (R32). Using this primer pair, only the full-length dystrophin transcript (product ‘a’) was amplified from untransfected cells or cells transfected with the control chimeraplast. However, an additional smaller PCR product (product ‘b’) was amplified from cells transfected with the targeting chimeraplast. Molecular weight markers are shown in the lane labeled ‘M’, and the lane labeled ‘C’ is a primer control lacking cDNA template. (C) Using different primer pairs, additional alternative spliced transcripts were identified following reverse transcription and nested PCR. Each product was purified from agarose gels and loaded individually in the lanes shown here. The outer and inner primers used in the nested PCR reactions that generated each product are shown above each lane, and the product designations (‘c’, ‘d’, ‘e’ and ‘f’; see Table 1) are shown at the bottoms of the lanes. (D) Direct sequencing of PCR products shown in panels (B) and (C) revealed that the product designated ‘a’ was indeed the full-length transcript with a normal intron 22/exon 23 junction and no skipping of exon 23. Sequence analysis of the smaller products (‘b’–‘f’) detected in MDX3-transfected cells revealed that these products represented alternatively spliced transcripts, each skipping exon 23 in addition to multiple exons upstream and downstream of the intron 22/exon 23 splice junction. The products resulted from the splicing of exons 21–31 (‘b’), exons 11–28 (‘c’), exons 17–31 (‘d’), exons 12–34 (‘e’) and exons 11–38 (‘f’). Sequences for products ‘c’ and ‘d’ were performed using reverse primers. Details of the product size, reading frame and predicted protein size for each of these are presented in Table 1.

 

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  		Table 1. Alternative splicing induced by MDX3
 
Because smaller transcripts are preferentially amplified by PCR, the intensity of bands on the gels does not reflect the relative abundance of the transcripts in the cell (see Fig. 6B). Normalizing the smaller products to the full-length transcript would thus overestimate the abundance of the alternatively spliced transcript. This is highlighted by the fact that the full-length product was always equally amplified when internal primers within 1–2 kb of each other (e.g. F19/R32 in Fig. 6B) were used in the nested PCR reaction. With internal primers increasingly distant from one another (e.g. F9/R43in or F9/R50in), only the alternatively spliced transcripts were amplified. This was not due to incomplete retrotranscription since the full-length transcript was always equally amplified when we used the F22/R32 primer pairs (data not shown). Therefore, this type of analysis does not provide data that accurately reflect the levels of the alternatively spliced transcripts relative to the full-length transcript.

The unexpected finding of multiple splice variants induced by a single base alteration at a single-splice junction led us to examine other splice junctions, specifically the donor and acceptor sites for exons that bracketed the skipped exons, for mutations. We did not expect this for two main reasons. First, there was no homology between the sequences of the other splice junctions and that of the intron 22/exon 23 junction. Thus, there would be no homologous pairing between MDX3 and any of those splice junctions that might lead to base changes. Second, MDX4 is nearly identical to MDX3 and yet did not induce any alternate splicing. Thus, it would be extraordinary if MDX3 were capable of inducing mutations at multiple, non-homologous splice sites along the dystrophin gene whereas MDX4, differing by only a single base from MDX3, had no activity whatsoever in inducing mutations. Still, based on the pattern of alternate splicing identified in MDX3-transfected cells (Table 1), we examined other splice sites for mutations. We PCR-amplified the donor splice site of the 5' exon and the acceptor splice site for the 3' exon for each novel splicing of distant exons to one another. In addition, we sequenced both the donor and acceptor splice sites for the first ‘skipped’ exon from either the 5' or 3' direction (see Fig. 7 for an example). All amplifications were taken through two rounds of PCR before sequencing, just as we did for the intron 22/exon 23 splice junction in Figure 2C. Figure 7 shows the sequence data for the case in which there was aberrant splicing between exon 17 and exon 31 (product ‘d’ from Fig. 6 and Table 1). There was no evidence of alterations in any bases in the consensus splice sequence or adjoining sequences for either site, even after two rounds of PCR amplification. Identical analysis was done for products ‘b’, ‘c’, ‘e’ and ‘f’ (Fig. 6 and Table 1), none of which showed any evidence of splice site mutations. Therefore, the only mutation identified in the dystrophin gene among these 25 splice sites was the single, targeted base at the intron 22/exon 23 boundary. These data suggest that mutations in single splice junctions can have more wide ranging effects than the skipping of a single exon in the splicing of a complex gene like dystrophin.



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  		Figure 7. Splice junction sequences. Genomic DNA was subjected to two rounds of PCR amplification using primers on either side of splice junctions of exons that were involved in the alternative splicing seen in MDX3-treated cells. To examine for splice site mutations that could account for aberrant splicing of exon 17 to exon 31, we sequenced the donor splice site of the exon 17/intron 17 boundary (‘donor’) and the acceptor splice site of the intron 30/exon 31 boundary (‘acceptor’). Because exons 18 and 30 were the first adjacent exons skipped on the 5' or 3' side, respectively, we also sequenced both the donor and acceptor splice sites of each of these exons as well (sites ‘a’, ‘b’, ‘c’ and ‘d’). No mutations were detected in any of the splice sites analyzed.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
We have extended the therapeutic potential of chimeraplast-mediated single base substitution from simply the correction of point mutations to the restoration of the reading frame by using the base conversion capability of chimeric oligonucleotides to alter RNA splicing. This approach would be applicable to the conversion of out-of-frame mutations (including large deletions) into in-frame mutations. In humans, this would have the potential to transform a severe DMD phenotype into a much milder BMD phenotype. A similar outcome is the goal of antisense-mediated exon skipping that has been applied to mutations in both the mouse and human dystrophin genes (8,10–12). While antisense-mediated exon skipping has shown promise, the use of chimeraplasts to induce exon skipping of the dystrophin gene would have several advantages. First, the corrections induced by chimeraplasts occur at the genomic level, thus avoiding the need to deliver oligonucleotides continuously or to express antisense constructs continuously in these cells in order to maintain dystrophin expression. Second, gene correction can occur not only in mature myofibers (21) but also in muscle precursor cells (20) that participate in muscle maintenance and repair. Thus, the therapeutic benefit would be stable not only in terms of correction at the genomic level, but also in terms of turnover in the tissue as muscle precursor cells fuse with existing myofibers or fuse with each other to form new myofibers. Currently, the efficiency of base conversion is at least an order of magnitude lower than would be necessary for therapeutic application, and this remains one of the major hurdles for this type of gene therapy technology.

Our results demonstrate that a targeting chimeraplast (MDX3) is capable of inducing a G-to-T conversion at the intron 22/exon 23 splice site of the dystrophin gene (Fig. 1B) and thereby disrupt the splicing of exon 22 to exon 23. However, this genomic conversion had more widespread effects on the splicing of the dystrophin pre-mRNA and resulted in the expression of multiple forms of the dystrophin protein, some with large internal deletions. Analysis of mRNA confirmed the presence of alternatively spliced transcripts with the number of excluded exons ranging from 11 to 27 (Fig. 6). Western blot analysis using an antibody specific for the C-terminal domain of the dystrophin protein consistently revealed the presence of at least three distinct protein products (Fig. 3A). Immunostaining using antibodies specific for different portions of the dystrophin protein confirmed that the dystrophin proteins produced had internal deletions (Fig. 3C).

The data presented led to the unexpected result that a mutation at a single base in one splice site could alter splicing more globally within the dystrophin gene. The chimeraplast sequence chosen for this experiment did not match any other genomic sequences reported in the BLAST database, thus ensuring the specificity of the oligonucleotides chosen for targeting. Altering even a single base in a chimeric oligonucleotide dramatically reduces its propensity to induce base conversions (23). Thus, there would be no reason to expect that a chimeraplast targeted to the intron 22/exon 23 junction would target a non-homologous splice junction. Along these same lines, it has been shown that chimeraplasts targeted for specific genomic sequences are capable on inducing single-base conversions in those genes but have no mutational propensity even in a gene with a highly homologous sequence (24). Thus, although we cannot totally exclude the possibility that MDX3 could have mutational activity elsewhere in the genome, it is highly unlikely that it is inducing base conversions specifically in splice sites in the dystrophin gene that have no homology to the targeted sequence. However, in order to support these theoretical arguments, we directly sequenced all of the affected splice sites and found evidence of base conversion only at the targeted base (Figs 2C and 7). Furthermore, as noted previously, any hypothesis that the alternate splicing induced by MDX3 is due to mutations of more sites than just the single base at the intron 22/exon 23 junction would have to take into account the fact that MDX4, identical to MDX3 other than a single base pair change, had no propensity to induce alternate splicing of the dystrophin gene.

Although MDX3 could theoretically have an effect on RNA processing directly (e.g. by an antisense-type activity), the possibility that the alternatively spiced products arose from persistent chimeraplasts acting on RNA processing is ruled out by our long-term results. The cells proliferate with a population doubling time of about 18 h. We see the same pattern of transcript and protein expression 3 months after transfection as we do within 1 week of transfection. At the 3-month time point, the amount of chimeraplast per cell would have decreased by a factor of 2120 from the time of transfection based purely on the dilution that would occur during cell division. This corresponds to a 1x1036-fold reduction in chimeraplast concentration and doesn't even take into account the degradation of chimeraplasts in the cells, which is likely to have at least as great an effect on the reduction of chimeraplast per cell as does cell division. Thus, the amount of chimeraplast in a cell at the 3-month time point would be so infinitesimal so as to be truly negligible. We therefore conclude that none of the results can be explained by persistent chimeraplast.

It is unclear why the alteration of a single splice site at the genomic level results in the expression of multiple splicing products, each skipping many exons both upstream and downstream of the targeted splice site. This phenomenon however has been observed in clinical cases of DMD patients in whom point mutations in splice sites lead to the skipping of multiple exons of the dystrophin gene (29,30). Altering a specific intron/exon sequence containing regulatory elements for the binding of splicing factors could disrupt the secondary structure necessary for coordinated splicing of multiple adjacent exons (31). The resulting mature RNA could thus display the skipping of multiple exons both upstream and downstream of the point mutation. Indeed, skipping of multiple contiguous exons has been observed when the splicing process is disrupted by the binding of antisense oligonucleotides to a single splice junction (9).

In the mdx mouse and in humans with DMD, there is a spontaneous appearance of myofibers that express dystrophin immunoreactivity (i.e. at least a portion of the dystrophin protein) (32). The numbers of these so-called ‘revertant’ fibers increase with age, and it is well established that the forms of the dystrophin protein expressed have internal deletions ranging from a few exons to many exons (33). The mechanism by which this occurs is unclear, but somatic mutations have been suggested (34). Recent studies have shown that revertant fibers within a single cluster all express the same form of dystrophin, suggesting that they are derived from a clonal expansion of a single muscle precursor cell whose progeny fuse with myofibers locally and confer on those fibers the ability to express internally deleted dystrophin proteins (33). Although immunohistochemical analysis indicates which exons are not expressed, these studies do not exclude the possibility that multiple transcripts, and thus multiple internally deleted forms of dystrophin, are expressed. Indeed, the variable intensity of staining using exon-specific antibodies is consistent with this possibility (33). Our results suggest that revertant fibers could arise from spontaneous, somatic mutations in splice site sequences. Such mutations could account for the variable pattern of transcripts and proteins that are expressed and the appearance of shortened forms of dystrophin in which multiple, contiguous exons are not expressed (33).

The systemic delivery of oligonucleotides remains a major hurdle for the clinical application of either antisense oligonucleotides or chimeraplasts to alter gene splicing (13). However, the finding that chimeraplast-mediated base conversion can lead to alternate splicing and the production of functional forms of dystrophin is encouraging for a therapeutic approach to DMD. The fact that multiple transcripts are produced and that several of these are in-frame demonstrate that the functionality of the combined protein products is not dependent on a single product being in-frame. Nevertheless, our findings demonstrate that much more work is needed in order to understand the mechanisms that regulate dystrophin gene splicing to be able to predict the effect of splice site alterations of such a large and complex gene as dystrophin. Understanding the mechanisms leading to multiple transcript production may shed light on the normal process of revertant fiber formation, a process that may also have therapeutic application if it can be understood and enhanced (35). Clearly, the modification of pre-mRNA splicing has great potential as a therapeutic approach to DMD and multiple technologies are advancing this novel therapeutic approach.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Mice
Mice of the mdx strain (C57BL/10ScSn-mdx) and the control C57 strain (C57BL/10SnJ) were obtained from the Jackson laboratory and were handled in accordance with guidelines of the Administrative Panel on Laboratory Animal Care of Stanford University.

Chimeraplast synthesis
Chimeric RNA/DNA oligonucleotides were synthesized as previously described (23) and were provided by Valigen Inc. (Newtown, PA, USA). The oligonucleotides were prepared with DNA and 2'-O-methyl RNA phosphoramidite nucleoside monomers on an Expedite Nucleic Aid Synthesizer (PerSeptive Biosystems, Framingham, MA, USA), purified by HPLC, and quantified by UV absorbance. More than 95% of the purified oligonucleotides were determined to be of full length. The targeting chimeraplast, designated MDX3 (Fig. 1A), was designed to induce a G-to-T conversion in the acceptor splice site of exon 23 of the dystrophin gene, thus destroying its consensus sequence (Fig. 1B). The control chimeraplast, designated MDX4 (Fig. 1A), differs from MDX3 by a single base and has no mismatches when aligned with the intron 22/exon 23 splice site. Thus, MDX4 should have all of the homologous pairing properties of MDX3 but none of its single base substitution activity in mdx cells.

Cell culture and transfection
Cells were derived from limb muscle of neonatal mdx and C57 mice as previously described (36). For growth, cells were plated on dishes coated with 5 µg/ml laminin (Invitrogen Life Technology, Carlsbad, CA, USA) and maintained in growth medium consisting of Ham's F10 nutrient mixture (Mediatech, Herndon, VA, USA) supplemented with 20% fetal bovine serum (Omega Scientific, Tarzana, CA, USA) and penicillin/streptomycin (Mediatech). Cell differentiation was induced by maintaining the cells in low serum medium (‘differentiation medium’) consisting of Dulbecco's modified Eagle's medium (Mediatech) supplemented with 2% horse serum (Invitrogen) and penicillin/streptomycin.

Myoblasts were plated in wells of six-well dishes (1x105 cells/well) 12 h prior to transfection. Chimeraplasts (10 µg) were complexed with 1 µl of Lipofectamine 2000 (GIBCO/BRL) for 30 min at room temperature in a total volume of 0.5 ml of Ham's F10. The complex was then added to the wells containing 1.5 ml of growth medium for 16 h. Transfection was stopped by replacing the transfection solution with fresh growth medium.

Western blot analysis
Cells were lysed in RIPA buffer (50 mM Tris–HCl, pH 7.4, 150 mM NaCl, 0.5% deoxycholate, and 1% Nonidet P-40) containing aprotinin (20 µg/ml), leupeptin (20 µg/ml), phenylmethylsulfonyl fluoride (10 µg/ml), and sodium orthovanadate (1 mM), and total protein in the extract was determined by the Bio-Rad protein assay (Bio-Rad, Hercules, CA, USA). For dystrophin detection, 350 µg of total protein from each sample were separated by electrophoresis (10 mA for 15 h) using 5% SDS–polyacrylamide gels, and then transferred (250 mA for 7 h) onto nitrocellulose membranes. Membranes were probed overnight at 4°C with an antibody directed toward the rod domain (Mandys-8, 1:400; Sigma) or the C-terminal domain (Dys-2, 1:50; Novacastra, Newcastle, UK) of the dystrophin protein as previously described (20). The membranes were blocked with 5% milk in PBS for 1 h at room temperature. Blots were washed with 0.05% Tween-20 in PBS and then reacted with a horseradish peroxidase-coupled anti-mouse secondary antibody (Amersham Pharmacia Biotech, Piscataway, NJ, USA). Specific antibody binding was detected using an enhanced chemiluminescent system (Amersham).

ARMS assay and restriction endonuclease analysis
Cultured myoblasts were rinsed twice with PBS and genomic DNA was extracted using the Wizard Genomic DNA extraction kit (Promega, Madison, WI, USA) according to the manufacturer's instructions. For each amplification reaction, 500 ng of total DNA extracted from myoblast cultures were digested for 2 h at 37°C with Sal I or NotI to unfold the supercoiled structure of the DNA as previously described (20).

The designations and sequences of all PCR primers are listed in Table 2. For the ARMS (amplification refractory mutation system) assay (37), we used a forward primer in intron 22 [F(int)22] and either of two reverse primers. One, designated ‘w’, was specific for the wild-type intron 22/exon 23 splice site sequence and the other, designated ‘m3’, was specific to the splice site sequence after a G-to-T transversion at nucleotide 3137 (see Fig. 1B). PCR settings and reaction conditions were as previously described (20). DNA products were fractionated on 2% agarose gels in Tris-acetate/EDTA buffer. PCR products were purified using Qiaquick PCR spin columns (Qiagen Inc., Valencia, CA, USA) and DNA sequencing was carried out using an Applied Biosystems ABI377 automated sequencer.


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  		Table 2. PCR primers
 
Restriction endonuclease analysis was performed on total genomic DNA. 200 ng of genomic DNA was subjected to PCR amplification using a forward primer in intron 22 [F(int)22] and a reverse primer in exon 23 (R23). Each amplification mixture contained 25 pmol of appropriate primer, 10% DMSO, 0.5 U Master Taq DNA polymerase (Takara, Panvera Corp., Madison, WI, USA), and 5 mM of each deoxyribonucleotide triphosphate. After an initial step of denaturation at 95°C for 5 min, amplification was performed for 35 cycles at 95°C for 1 min followed by annealing at 55°C for 3 min and extension at 72°C for 2 min. Amplification reactions were terminated by an additional extension step at 72°C for 10 min. PCR products were loaded on a 2% agarose gel and the expected 150 bp amplification product was excised from the gel and purified using Qiagen gel extraction kit. Of the 30 µl of the resuspended DNA product, 5 µl were subjected to a second round of PCR using the same conditions previously described with the addition of 10 mCi [32P]dCTP. The radioactive samples were purified using Amicon Microcon®-PCR Centrifugal Filter Devices (Millipore Corporation, Bedford, MA, USA) and resuspended in 50 µl of H20. Ten microliters were digested for 1 h at 37°C with Hinf I restriction enzyme and loaded on an 8% non-denaturing acrylamide gel. Bands were visualized after direct exposure of the gel to X-ray film.

Nested RT–PCR
Total RNA was extracted from cultured myotubes using TRI-REAGENT (Sigma). For each reaction, 100 ng of RNA were treated with 1 U of DNase I (GIBCO/BRL) at room temperature and retrotranscribed using One-Step RT–PCR (Qiagen). Reverse transcription and first strand amplification were carried out using a reverse primer in exon 33 (R33) paired with a forward primer in exon 22 (F22). Nested PCR was carried out on 2 µl of first strand amplification reaction using the forward (inner) primer in exon 22 (F22in) and reverse primers in exons 23 (R23), 24 (R24), 25 (R25), 30 (R30) and 32 (R32).

To detect alternative splicing products outside the region between exons 22 and 32 (see Fig. 6), a set of different reverse transcription and first strand amplification reactions were carried out using reverse primers in exons 33 (R33), 35 (R35out), 43 (R43out) and 50 (R50out) paired with a forward primer in exon 8 (F8). Nested PCR was performed with forward primers in exons 9 (F9) and 19 (F19) paired with reverse primers in exons 32 (R32), 35 (R35in), 43 (R43in) and 50 (R50in). Amplification was carried out for 25 cycles at 94°C for 30 s, 55°C for 2 min and 72°C for 2 min. PCR products were excised from the agarose gel, purified using a Qiagen gel extraction kit, and sequenced using an automated sequencer.

Splice site sequence analysis
Sequences of the mouse dystrophin gene were retrieved from the NCBI mouse genome database. Genomic DNA was isolated as previously described from untransfected cells or cells transfected with MDX3 that were kept in culture for 3 months after transfection. Analyses of splice site sequences were carried out using 100 ng of genomic DNA. The intron 22/exon 23 splice junction was amplified using the F(int)22 and R23 primers previously described. For all other splice junctions, forward and reverse primers were designed to amplify ~400 bp, including the whole exon and both the donor and acceptor splice sites. Amplification was carried out for 30 cycles using the settings previously described for the restriction endonuclease analysis. PCR products were resolved on agarose gels, purified and resuspended in 30 µl of H2O. A second round of amplification was carried out using 2 µl of the purified product for an additional 30 cycles. Second round PCR products were then purified and sequenced as previously described. Primer sequences and electropherograms are available upon request.

Immunohistochemistry
Immunostaining was performed as previously described (20,36). Briefly, cells cultured on coverslips were fixed in ice-cold ethanol for 10 min, rinsed in PBS, and permeabilized in 0.3% Triton X-100 for 10 min at room temperature. A solution containing 5% heat-inactivated normal goat serum and 1 mg/ml BSA in PBS was used as blocking solution for 30 min. Cells were then incubated with primary antibodies to dystrophin (Mandys-18, 1:100, generous gift of Dr Glenn Morris, The North East Wales Institute, Wrexham, UK; Mandys-8, 1:200, Novacastra; Dys-2, 1:25, Novacastra) or {alpha}-dystroglycan (VIA4-1, 1:100; Upstate Biotechnology, Lake Placid, NY, USA) overnight at 4°C. Cells were washed in PBS before incubation for 1 h at room temperature with an AlexaTM 546-coupled goat–anti-mouse (H+L) (1:250; Molecular Probes, Eugene, OR, USA) or FITC-conjugated-goat-anti-mouse (whole molecule; 1:200; Cappel ICN Pharmaceuticals Inc., Aurora, OH, USA) secondary antibodies. Coverslips were mounted using Vecta Shield (Vector Inc., Burlingame, CA, USA) for fluorescence microscopy.


   ACKNOWLEDGEMENTS
 
The authors would like to thank Dr Glenn Morris, The North East Wales Institute, Wrexham, UK, for generously providing exon-specific antibodies to dystrophin. The authors would also like to thank Dr Steve Wilton for helpful discussions. This work was supported by a grant from Duchenne Parent Project/NL and Italia and Aktion Benni & Co. e.V., Germany to C.B. and by a grant from the Muscular Dystrophy Association (USA) to T.A.R.


   FOOTNOTES
 
* To whom correspondence should be addressed at: Department of Neurology and Neurological Sciences, Stanford University Medical Center, Room A-343, Stanford, CA 94305-5235, USA. Tel: +1 6508583976; Fax: +1 6508583935; Email: rando{at}stanford.edu Back


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 INTRODUCTION
 RESULTS
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