<p>Duchenne muscular dystrophy (DMD) is a severe hereditary disorder caused by a loss of dystrophin, an essential musculoskeletal protein. Decades of promising research have yielded only modest gains in survival and quality of life for these patients and there have been no approved gene therapies for DMD to date. There are two significant hurdles to creating effective gene therapies for DMD; it is difficult to deliver a replacement dystrophin gene due to its large size and current strategies to restore the native dystrophin gene likely require life-long administration of a gene-modifying drug. This thesis presents a novel method to address these challenges through restoring dystrophin expression by genetically correcting the native dystrophin gene using engineered nucleases that target one or more exons in a mutational hotspot in exons 45-55 of the dystrophin gene. Importantly, this hotspot mutational region collectively represents approximately 62% of all DMD mutations. In this work, we utilize various engineered nuclease platforms to create genetic modifications that can correct a variety of DMD patient mutations.</p><p>Initially, we demonstrate that genome editing can efficiently correct the dystrophin reading frame and restore protein expression by introducing micro-frameshifts in exon 51, which is adjacent to a hotspot mutational region in the dystrophin gene. Transcription activator-like effector nucleases (TALENs) were engineered to mediate highly efficient gene editing after introducing a single TALEN pair targeted to exon 51 of the dystrophin gene. This led to restoration of dystrophin protein expression in cells from DMD patients, including skeletal myoblasts and dermal fibroblasts that were reprogrammed to the myogenic lineage by MyoD. We show that our engineered TALENs have minimal cytotoxicity and exome sequencing of cells with targeted modifications of the dystrophin locus showed no TALEN-mediated off-target changes to the protein coding regions of the genome, as predicted by in silico target site analysis. </p><p>In an alternative approach, we capitalized on the recent advances in genome editing to generate permanent exclusion of exons by using zinc-finger nucleases (ZFNs) to selectively remove sequences important in specific exon recognition. This strategy has the advantage of creating predictable frame restoration and protein expression, although it relies on simultaneous nuclease activity to generate genomic deletions. ZFNs were designed to remove essential splicing sequences in exon 51 of the dystrophin gene and thereby exclude exon 51 from the resulting dystrophin transcript, a method that can potentially restore the dystrophin reading frame in up to 13% of DMD patients. Nucleases were assembled by extended modular assembly and context-dependent assembly methods and screened for activity in human cells. Selected ZFNs had moderate observable cytotoxicity and one ZFN showed off-target activity at two chromosomal loci. Two active ZFN pairs flanking the exon 51 splice acceptor site were transfected into DMD patient cells and a clonal population was isolated with this region deleted from the genome. Deletion of the genomic sequence containing the splice acceptor resulted in the loss of exon 51 from the dystrophin mRNA transcript and restoration of dystrophin expression in vitro. Furthermore, transplantation of corrected cells into the hind limb of immunodeficient mice resulted in efficient human dystrophin expression localized to the sarcolemma. </p><p>Finally, we exploited the increased versatility, efficiency, and multiplexing capabilities of the CRISPR/Cas9 system to enable a variety of otherwise challenging gene correction strategies for DMD. Single or multiplexed sgRNAs were designed to restore the dystrophin reading frame by targeting the mutational hotspot at exons 45-55 and introducing either intraexonic small insertions and deletions, or large deletions of one or more exons. Significantly, we generated a large deletion of 336 kb across the entire exon 45-55 region that is applicable to correction of approximately 62% of DMD patient mutations. We show that, for selected sgRNAs, CRISPR/Cas9 gene editing displays minimal cytotoxicity and limited aberrant mutagenesis at off-target chromosomal loci. Following treatment with Cas9 nuclease and one or more sgRNAs, dystrophin expression was restored in Duchenne patient muscle cells in vitro. Human dystrophin was detected in vivo following transplantation of genetically corrected patient cells into immunodeficient mice. </p><p>In summary, the objective of this work was to develop methods to genetically correct the native dystrophin as a potential therapy for DMD. These studies integrate the rapid advances in gene editing technologies to create targeted frameshifts that restore the dystrophin gene around patient mutations in non-essential coding regions. Collectively, this thesis presents several gene editing methods that can correct patient mutations by modification of specific exons or by deletion of one or more exons that results in restoration of the dystrophin reading frame. Importantly, the gene correction methods described here are compatible with leading cell-based therapies and in vivo gene delivery strategies for DMD, providing an avenue towards a cure for this devastating disease.</p> / Dissertation

Our previous studies have demonstrated that RNA can serve as a template for double-strand break (DSB) repair in the yeast Saccharomyces cerevisiae using synthetic RNA-containing oligonucleotides (oligos). Following this initial work, we show that the RNA tract of RNA-containing oligos can be copied into DNA to transfer a genetic change at the chromosomal level also in the bacterium Escherichia coli and in human cells. Exploiting the use of oligos containing ribonucleoside monophosphates (rNMPs), we developed a molecular approach to generate RNA/DNA hybrids of chosen sequence and structure at the chromosomal level in both yeast and E. coli cells. Such technique allows us to study how rNMPs present in the DNA genome of cells are tolerated by cells, what factors recognize and target rNMPs in DNA and to what extent the embedded rNMPs may alter genome integrity. Here we proved that mispaired rNMPs embedded into genomic DNA, if not removed, serve as templates for DNA synthesis during chromosomal replication and produce a genetic change. We discovered that mispaired rNMPs that are embedded in genomic DNA are not only targeted by ribonucleases H (RNases H) but also by the mismatch repair (MMR) system both in yeast and in E. coli. Our data reveal novel substrates for the MMR system, and also uncover an unpredicted competition between RNase H and MMR for the RNA/DNA mispairs.

No description available.

Here we developed multiple genetic systems through which genetic modifications driven by DNA breaks caused by the I-SceI nuclease can be assayed in the yeast Saccharomyces cerevisiae and in human cells. Using the delitto perfetto approach for site-directed mutagenesis in yeast, we generated isogenic strains in which we could directly compare the recombination potential of different I-SceI variants. By genetic engineering procedures, we generated constructs in human cells for testing the recombination activity of the same I-SceI variants. Both in yeast and human cells we performed gene correction experiments using oligonucleotides (oligos) following modification and/or optimization of existing gene targeting protocols and development of new ones. We demonstrated that an I-SceI nicking enzyme can stimulate recombination on the chromosome in S. cerevisiae at multiple genomic loci. We also demonstrated in yeast that an I-SceI-driven nick can activate recombination 10 kb distant from the initial site of the chromosomal lesion. Moreover we demonstrated that an I-SceI nick can stimulate recombination at the site of the nick at episomal and chromosomal loci in human cells. We showed that an I-SceI double-strand break (DSB) could trigger recombination up to 2 kb distant from the break at an episomal target locus in human cells, though the same was not observed for the nick. Overall, we demonstrated the capacity for I-SceI nick-induced recombination in yeast and human cells. Importantly, our findings reveal that the nick stimulates gene correction by oligos differently from a DSB lesion, as determined by genetic and molecular analyses in yeast and human cells. This research illustrates the promise of targeted gene correction following generation of a nick.

Rare diseases, when considered as a whole, affect up to 7% of the population, which would represent 3.5 million individuals in the United Kingdom alone. However, while 'personalised medicine' is now yielding remarkable results using recent sequencing technologies in terms of diagnosing genetic conditions, we have made much less headway in translating this patient information into therapies and effective treatments. Even with recent calls for greater research into personalised treatments for those affected by a rare disease, progress in this area is still severely lacking, in part due to the astronomical cost and time involved in bringing treatments to the clinic. Gene correction using the recently-described genome editing technology CRISPR/Cas9, which allows precise editing of DNA, offers an exciting new avenue of treatment, if not cure, for rare diseases; up to 80% of which have a genetic component. This system allows the researcher to target any locus in the genome for cleavage with a short guide-RNA, as long as it precedes a highly ubiquitous NGG sequence motif. If a repair sequence is then also provided, such as a wild-type copy of the mutated gene, it can be incorporated by homology-directed repair (HDR), leading to gene correction. As both guide-RNA and repair template are easily generated, whilst the machinery for editing and delivery remain the same, this system could usher in the era of 'personalised medicine' and offer hope to those with rare genetic diseases. However, currently it is difficult to test the efficacy of CRISPR/Cas9 for gene correction, especially in vivo. Therefore, in my PhD I have developed a novel fluorescent reporter system which provides a rapid, visual read-out of both non-homologous end joining (NHEJ) and homology-directed repair (HDR) driven by CRISPR/Cas9. This system is built upon a cassette which is stably and heterozygously integrated into a ubiquitously expressed locus in the mouse genome. This cassette contains a strong hybrid promoter driving expression of membrane-tagged tdTomato, followed by a strong stop sequence, and then membrane-tagged EGFP. Unedited, this system drives strong expression of membrane-tdTomato in all cell types in the embryo and adult mouse. However, following the addition of CRISPR/Cas9 components, and upon cleavage, the tdTomato is rapidly excised, resulting via NHEJ either in cells without fluorescence (due to imperfect deletions) or with membrane-EGFP. If a repair template containing nuclear tagged-EGFP is also supplied, the editing machinery may then use the precise HDR pathway, which results in a rapid transition from membrane-tdTomato to nuclear- EGFP. Thereby this system allows the kinetics of editing to be visualised in real time and allows simple scoring of the proportion of cells which have been edited by NHEJ or corrected by HDR. It therefore provides a simple, fast and scalable manner to optimise reagents and protocols for gene correction by CRISPR/Cas9, especially compared to sequencing approaches, and will prove broadly useful to many researchers in the field. Further to this, I have shown that methods which lead to gene correction in our reporter system are also able to partially repair mutations found in the disease-causing gene, Zmynd10; which is implicated in the respiratory disorder primary ciliary dyskinesia (PCD), for which there is no effective treatment. PCD is an autosomal-recessive rare disorder affecting motile cilia (MIM:244400), which results in impaired mucociliary clearance leading to neonatal respiratory distress and recurrent airway infections, often progressing to lung failure. Clinically, PCD is a chronic airway disease, similar to CF, with progressive deterioration of lung function and lower airway bacterial colonization. However, unlike CF which is monogenic, over 40 genes are known to cause PCD. The high genetic heterogeneity of this rare disease makes it well suited to such a genome editing strategy, which can be tailored for the correction of any mutated locus.