<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

Engineered DNA-binding proteins enable targeted manipulation of the genome. Zinc fingers are the most well characterized DNA-binding domain and for many years research has focused on understanding and manipulating the sequence-specificities of these proteins. Recently, major advances in the ability to engineer zinc finger proteins, as well as the discovery of a new class of DNA-binding domains - transcription activator-like effectors (TALEs), have made it possible to rapidly and reliably engineer proteins targeted to any sequence of interest. With this capability, focus has shifted to exploring the applications of this powerful technology. In this dissertation I explore three important applications of engineered DNA-binding proteins.

Die einzellige Grünalge Chlamydomonas reinhardtii ist ein vielseitiger Modellorganismus sowohl in der Grundlagenforschung als auch für biotechnologische Anwendung. Für die genetische Veränderung wurden verschiedene Methoden entwickelt, jedoch ist die gezielte Modifikation kerncodierter Gene immernoch sehr schwierig. In dieser Arbeit wird eine Strategie vorgestellt, die es ermöglicht, kerncodierte Gene in Chlamydomonas gezielt mit sequenzspezifischen Zinkfinger-Nukleasen zu verändern. Das COP3-Gen, welches den lichtaktivierbaren Ionenkanal Kanalrhodopsin-1 codiert, diente hierbei als Zielsequenz der für die Deletion hergestellten Zinkfinger-Nukleasen. Um eine Charakterisierung der ZFNs zu ermöglichen, wurde ein Modelstamm generiert, der ein inaktiviertes Markergen enthält. Die Inaktiverung erfolgte hierbei durch Insertion der COP3-ZFN Zielsequenz. Die Transformation dieses Modellstamms mit ZFN codierender Plasmid-DNA und einem Reparatur-Template ermöglichte die Wiederherstellung der Markeraktivität und eine Selektion Antibiotika-resistenter Kolonien. Wenn in diesen Experimenten zusätzlich ein COP3 veränderndes Template benutzt wurde, enthielt 1% der analysierten Klone ein mutiertes COP3-Gen. Der Chlamydomonas Augenfleck ist ein lichtsensitives Organell mit entscheidender Funktion für die phototaktische Orientierung der Alge. Eine Deletionsmutante des Blaulicht-Photorezeptors Phototropin zeigte in Experimenten eine veränderte Regulation der lichtabhängigen Augenfleckgröße. Durch Komplementierung der Phototropin-Dysfunktion konnte der lichtabhängige Regulationsprozess wiederhergestellt werden. Die Expression der Phototropin-Kinasedomäne führte zu einer lichtunabhängigen Reduktion der Augenfleckfläche. Interessanterweise führte auch die Expression der N-terminalen LOV-Domänen zu einer geänderten Regulation des Augenflecks und der Phototaxis. Dies deutet, zusätzlich zur Lichtregulation der Kinasedomäne, auf eine zelluläre Signalfunktion der LOV-Domänen hin. / The unicellular green alga Chlamydomonas reinhardtii is a versatile model for fundamental and biotechnological research. A wide toolset for genetic manipulation has been developed for this alga, but specific modification of nuclear genes is still not routinely possible. Here we present a nuclear gene targeting strategy for Chlamydomonas that is based on the application of zinc-finger nucleases (ZFNs). Initially, we designed a set of ZFNs for targeting the COP3 gene that encodes the light-activated ion channel channelrhodopsin-1. To evaluate the designed ZFNs, we constructed a model strain by inserting a non-functional selection marker interspaced with a short COP3 target sequence into the nuclear genome. Upon co-transformation of this recipient strain with the engineered ZFNs and a DNA repair template, we were able to restore marker activity and select antibiotic resistant clones with active nucleases. In cases where cells were co-transformed with a modified COP3 template, 1% of these clones contained a modified COP3 locus as well. The eyespot of Chlamydomonas is a light-sensitive organelle important for phototactic orientation of the alga. Here we found that eyespot size is downregulated in light. In a strain in which the blue light photoreceptor phototropin was deleted, the light regulation of the eyespot size was affected. We restored this dysfunction in different phototropin complementation experiments. Complementation with the phototropin kinase fragment reduced the eyespot size, independent of light. Interestingly, overexpression of the N-terminal LOV-domains alone also affected eyespot size and phototaxis, suggesting that aside from activation of the kinase domain, they fulfill an independent signaling function in the cell. We propose that phototropin is a light regulator of phototaxis that desensitizes the eyespot when blue light intensities increase.

Die Modifikation von Genen ist in den molekularen Biowissenschaften ein fundamentales Werkzeug, um die Funktion von Genen zu studieren (Reverse Genetik). Diese Arbeit hat erfolgreich Zinkfinger- und CRISPR/Cas9-Nukleasen für die Verwendung in C. reinhardtii etabliert, um Gene im Kerngenom gezielt auszuschalten und präzise zu verändern. Basierend auf vorausgegangener Arbeit mit Zinkfingernukleasen (ZFN) konnte die Transformationseffizienz um das 300-fache verbessert werden, was die Inaktivierung von Genen auch in motilen Wildtyp-Zellen ermöglichte. Damit war es möglich, die Gene für das Kanalrhodopsin-1 (ChR1), Kanalrhodopsin-2 (ChR2) und das Chlamyopsin-1/2-Gen (COP1/2) einzeln und gemeinsam auszuschalten. Eine Analyse der Phototaxis in diesen Stämmen ergab, dass die Phototaxis durch Inaktivierung von ChR1 stärker beeinträchtigt ist als durch Inaktivierung von ChR2. Um das CRISPR/Cas9-System zu verwenden, wurden die Transformationsbedingungen so angepasst und optimiert, dass der Cas9-gRNA-Komplex als in vitro hergestelltes Ribonukleoprotein in die Zellen transformiert wurde. Um die Bedingungen für präzise Genmodifikationen zu messen und zu verbessern, wurde das SNRK2.2-Gen als Reportergen für eine „Blau-Grün Test“ etabliert. Kleine Insertionen von bis zu 30 bp konnten mit kurzen Oligonukleotiden eingefügt werden, während größere Reportergene (mVenus, SNAP-Tag) mithilfe eines Donor-Plasmids generiert wurden. In dieser Arbeit konnten mehr als 20 nicht-selektierbare Gene – darunter 10 der 15 potenziellen Photorezeptorgene – mit einer durchschnittlichen Mutationsrate von 12,1 % inaktiviert werden. Insgesamt zeigt diese Arbeit in umfassender Weise, wie Gen-Inaktivierungen und Modifikationen mithilfe von ZFNs und des CRISPR/Cas9-Systems in der Grünalge C. reinhardtii durchgeführt werden können. Außerdem bietet die Sammlung der zehn Photorezeptor-Knockouts eine aussichtsreiche Grundlage, um die Vielfalt der Photorezeptoren in C. reinhardtii zu erforschen. / Gene editing is a fundamental tool in molecular biosciences in order to study the function of genes (reverse genetics). This study established zinc-finger and CRISPR/Cas9 nucleases for gene editing to target and inactivate the photoreceptor genes in C. reinhardtii. In continuation of previous work with designer zinc-finger nucleases (ZFN), the transformation efficiency could be improved 300-fold, which enabled the inactivation of genes in motile wild type cells. This made it possible to disrupt the Channelrhodopsin-1 (ChR1), Channelrhodopsin-2 (ChR2) and Chlamyopsin-1/2 (COP1/2) genes individually and in parallel. Phototaxis experiments in these strains revealed that the inactivation of ChR1 had a greater effect on phototaxis than the inactivation of ChR2. To apply the CRISPR/Cas9 system, the transformation conditions were adapted and optimized so that the Cas9-gRNA complex was successfully electroporated into the cells as an in vitro synthesized ribonucleoprotein. This approach enabled gene inactivations with CRISPR/Cas9 in C. reinhardtii. In order to measure and improve the conditions for precise gene modifications, the SNRK2.2 gene was established as a reporter gene for a ‘Blue-Green test’. Small insertions of up to 30 bp were inserted using short oligonucleotides, while larger reporter genes (mVenus, SNAP-tag) were integrated using donor plasmids. Throughout this study, more than 20 non-selectable genes were disrupted, including 10 of the photoreceptor genes, with an average mutation rate of 12,1 %. Overall, this work shows in a comprehensive way how gene inactivations and modifications can be performed in green alga C. reinhardtii using ZFNs or CRISPR/Cas9. In addition, the collection of the ten photoreceptor knockouts provides a promising source to investigate the diversity of photoreceptor genes in C. reinhardtii.