The causative role of the huntingtin (HTT) gene in Huntington's disease (HD) has been identified more than 25 years ago. The extension of CAG repeat stretch over 39 repeats in exon 1 of one HTT allele results in full penetrance of this neurodegenerative disorder. While the identification of the causative mutation raised hopes that development of the therapeutic compound will be easily achievable, the patients and their families are still waiting for treatment until now. The main reason for that might be the complex cellular function HTT that makes the determination of the pathologic mechanism difficult and the development of treatments even more challenging. Although a lot of different animal models have been generated until now, establishing a suitable model has still not been achieved yet. Due to its anatomy, physiology, and genetics, the minipig seems to be a suitable candidate for neurodegenerative disease models. Indeed, the existing Transgenic (Tg) Libechov minipig model manifests signs typical for HD in patients, but on the other hand significant inconsistencies have also been observed. The finding of malformation that partially shows the situation in human patients is true for both, the male reproductive tract as well as for the brain. The reason for this might be the fact the genetic...

In this thesis, I argue that the treatment/enhancement distinction that is central to the ethical debate concerning germline genome editing and CRISPR is too vague to be ethically and normatively guiding. The problem of vagueness is twofold, being both a semantic and epistemic issue. This vagueness creates borderline cases, cases that cannot be properly defined as either treatment or enhancement, I call this The Borderline Cases Argument. These borderline cases enable a slippery slope towards eugenic practices, radical enhancement and dangerous applications of CRISPR. The distinction therefore fails to be action guiding as it cannot distinguish treatment from enhancement as well as failing to correspond to what is genuinely morally problematic with germline genome editing and not, I call this The Argument of Missing the Point. In using the treatment/enhancement distinction we therefore risk losing control over how CRISPR is used and for what purposes.

One of the major challenges facing medicine and drug discovery is the large number of genetic diseases caused by inherited mutations leading to a toxic gain-of-function, or loss-of-function of the disease protein. Microbiology offered a new glimpse of hope to address those disorders with the adaptation of the bacterial CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats) defense system as a genome editing tool. Cas9 is a unique CRISPR-associated endonuclease protein that can be easily programmed with an RNA [a single-guide RNA (sgRNA)] that is complementary to nearly any DNA locus. Cas9 creates a double-stranded break (DSB) that can be exploited to knock out toxic genes or replenish therapeutic expression levels of essential proteins. In addition to a matching sgRNA sequence, Cas9 requires the presence of a short signature sequence [a protospacer adjacent motif (PAM)] flanking the target locus. Over the past few years, several Cas9-based therapeutic platforms have emerged to correct DNA mutations in a wide range of mammalian cell lines, ex vivo, and in vivo by adapting recombinant adeno-associated virus (rAAV). However, most of the applications of Cas9 in the field have been limited to Streptococcus pyogenes (SpyCas9), which, in its wild-type form, suffers from inaccurate editing at off-target sites. It is also difficult to deliver via an all-in-one (sgRNA+Cas9) rAAV approach due to its large size. In this thesis, I describe other Cas9 nucleases and their development as new AAV-based genome editing platforms for therapeutic editing in vivo in mouse disease models. In the first part of this thesis, I develop the all-in-one AAV strategy to deliver a Neisseria meningitidis Cas9 ortholog (Nme1Cas9) in mice to reduce the level of circulating cholesterol in blood. I also help characterize an enhanced Cas9 from another meningococcus strain (Nme2Cas9) and show that it is effective in performing editing not only in mammalian cell culture, but also in vivo by all-in-one AAV delivery. Additionally, I describe two AAV platforms that enable advanced editing modalities in vivo: 1) segmental DNA deletion by delivering two sgRNAs (along with Nme2Cas9) in one AAV, and 2) precise HDR-based repair by fitting Nme2Cas9, sgRNA and donor DNA within a single AAV capsid. Using these tools, we successfully treat two genetic disorders in mice, underscoring the importance of this powerful duo of AAV and Cas9 in gene therapy to advance novel treatment. Finally, I present preliminary data on how to use these AAV.Nme2Cas9 vectors to treat Alexander Disease, a rare progressive neurological disorder. These findings provide a platform for future application of gene editing in therapeutics.

京都大学 / 新制・課程博士 / 博士(医学) / 甲第24485号 / 医博第4927号 / 新制||医||1063(附属図書館) / 京都大学大学院医学研究科医学専攻 / (主査)教授 江藤 浩之, 教授 湊谷 謙司, 教授 大鶴 繁 / 学位規則第4条第1項該当 / Doctor of Medical Science / Kyoto University / DGAM

<p>  Non-model microorganisms offer a wealth of biotechnological potential that may be leveraged to address a variety of global grand challenges. These include challenges in carrying out complex or altogether new chemistries, discovery and production of bioactive molecules, sustainable production of biochemicals and bioproducts from renewable feedstocks, and improving agricultural practices for responsible management of carbon. Specifically, using renewable plant biomass as a substrate for production of fuels and or chemicals offers a near ubiquitous supply that does not compete with food or petrochemicals. Alternatively, identifying new natural products will be essential to addressing the ever-increasing occurrence of antibiotic resistance. Non-model organisms may provide elegant solutions to many of these challenges, whether by possessing new or more efficient strategies to depolymerize lignocellulose, by encoding enzymes with increased stabilities and or specific activities, or perhaps by containing rich biosynthetic capabilities for production of previously unidentified natural products, among others. Yet efforts to leverage non-model microorganisms for their diverse biotechnological potential remain limited to a variety of often difficult, yet not insurmountable challenges.</p> <p>     In this work, I propose anaerobic gut fungi (Neocallimastigomycota) as a robust microbial system that may be leveraged to efficiently depolymerize crude lignocellulose, increase animal nutrition, or identify novel natural products. To this end, I detail the first chromosomally resolved genome assembly of anaerobic fungi (<em>Piromyces communis </em>var. <em>indianae</em> UH3-1). I investigate the genome organization of this isolate and describe how acquisition of Carbohydrate Active EnZymes (CAZymes) contribute to the robust lignocellulolytic activity of gut fungi. I then detail efforts to build a nascent genetic engineering toolbox for these anaerobic organisms. With the acquisition of the first chromosomally resolved genome assemblies, I identify a basic set of genetic parts needed for a genetic engineering toolkit. I show these parts are functional and detail methods to enable higher throughput testing in vivo. I subsequently detail efforts to construct the first preliminary CRISPR tools for anaerobic fungi as these will be essential to establish precise DNA targeting in future strain engineering efforts. I then describe the role of epigenetics in anaerobic fungi, detailing the extent to which it may be leveraged to control gene expression. Finally, I provide a discussion of this work and describe how it may guide future efforts to domesticate these organisms. Collectively, this work provides the first chromosomally resolved genome assembly as a resource for the community, along with genetic tools and techniques to begin domesticating these non-model organisms. Importantly, this work reveals that despite the challenges associated with anaerobic microbes of relatively high complexity, they are not insurmountable, and thus efforts to domesticate them are feasible.</p>

Human Immunodeficiency Virus (HIV) infection initiates major metabolic and cell- survival complications. Anti-retroviral therapy (ART) is the current approach to suppress active HIV replication to a level of undetected viral load, but it is not a curative approach. Newer and sophisticated gene editing technologies could indeed be a potent antiviral therapy to achieve a clinical sterilization/cure of HIV infection. Chronic HIV patients, even under a successful ART regimen, exhibit a low-grade inflammation, immune senescence, premature aging, telomeric DNA attrition, T cell apoptosis, and cellular homeostasis. In this dissertation, we investigated CD4 T cell homeostasis, degree of T cell apoptosis, an associated telomeric DNA damage, DNA damage repair signaling, and the apoptotic pathways in CD4 T cells during HIV infection with or without ART treatment. Our data support a DNA damage accumulation, and impaired DNA damage repair in chromosome ends via recruitment of 53BP1 protein to the damaged foci. We found that a key player of DNA damage and repair enzyme, ATM, and its associated checkpoint proteins (CHK1, CKH2) are affected by HIV infection. HIV infection also altered another multifunctional master regulator protein AKT that is crucial in maintaining cellular homeostasis. Curing HIV is the ultimate redemption against HIV-associated complications. To explore the possibility of a functional cure, we investigated the use of a transient and a non-viral CRISPR/Cas9-based gene editing technology targeting the latently incorporated HIV provirus. After performing a nucleofection/electroporation using an in vitro formulated ribonucleoprotein (RNP) constituting a synthetic guide RNA (gRNA) and Cas9 nuclease protein, we demonstrated a significant (maximum 97%) reduction of HIV-mRNA and p24-capsid protein expression, upon stimulation (using PMA) and latency reactivation of latently HIV-infected CD4 T cells and latent-monocytes. Notably, the RNP treatment did not induce any cytotoxic effects, without affecting the abilility of cell proliferation. A sequence specific cleavage of HIV-provirus in two crucial gene locations (targeting vpr/tat genes) showed the most significant suppression of HIV reactivation or latency reversal. We have used DNA sequencing, and T7EI assay to confirm the target-site-specific cleavage of the HIV-proviral genome. Our data confirm the activation of non- homologous end joining (NHEJ) pathway to repair the double-stranded DNA break created by the CRISPR/Cas9 treatment. Taken together, this study provides a new gene therapeutic approach using synthetic gRNA/Cas9 targeting HIV genome, which warrant further in vivo animal and human studies.

Tyrosine site-specific recombinases (Y-SSRs) are DNA editing enzymes that play a valuable role for the manipulation of genomes, due to their precision and versatility. They have been widely used in biotechnology and molecular biology for various applications, and are slowly finding their spot in gene therapy in recent years. However, the limited number of available Y-SSR systems and their often narrow target specificity have hindered the full potential of these enzymes for advanced genome engineering. In this PhD thesis, I conducted a comprehensive investigation of novel Y-SSRs and their potential for advancing genome engineering. This PhD thesis aims to address the current limitations in the genetic toolbox by identifying and characterizing novel Cre-type recombinases and demonstrating their impact on the directed evolution of designer recombinases for precise genome surgery. To achieve these aims, I developed in a collaboration a comprehensive prediction pipeline, combining a rational bioinformatical approach with knowledge of the biological functions of recombinases, to enable high success rate and high-throughput identification of novel tyrosine site-specific recombinase (Y-SSR) systems. Eight putative candidates were molecularly characterized in-depth to ensure their successful integration into future genome engineering applications. I assessed their activity in prokaryotes (E. coli) and eukaryotes (human cell lines), and determined their specificity in the sequence space of all known Cre- type target sites. The potential cytotoxicity associated with cryptic genomic recombination sites was also explored in the context of recombinase applicability. This approach allowed the identification of novel Y-SSRs with distinct target sites, enabling simultaneous use of multiple Y-SSR systems, and provided knowledge that will facilitate the assignment of novel and known recombinases to specific uses or organisms, ensuring their safe and effective implementation. The introduction of these novel Y-SSRs into the genome engineering toolbox opens up new possibilities for precise genome manipulation in various applications. The broader targetability offered by these enzymes could accelerate the development of novel gene therapies, as well as advance the understanding of gene function and regulation. Moreover, these recombinases could be used to design custom genetic circuits for synthetic biology, allowing researchers to create more complex and sophisticated cellular systems. Finally, I introduced the novel Y-SSRs into efforts aimed at developing designer recombinases for precise genome surgery, demonstrating their impact on accelerating the directed evolution process. Therapeutically relevant recombinases with altered DNA specificity have been developed for excision or inversion of specific DNA sequences. However, the potential for evolving recombinases capable of integrating large DNA cargos into naturally occurring lox-like sites in the human genome remained untapped so far. Thus, I embarked on evolving the Vika recombinase to mediate the integration of DNA cargo into a native human sequence. I discovered that Vika could integrate DNA into the voxH9 site in the human genome, and then, I enhanced the process through directed evolution. The evolved variants of Vika displayed a marked improvement in integration efficiency in bacterial systems. However, the translation of these results into mammalian systems has not yet been entirely successful. Despite this, the study laid the groundwork for future research to optimize the efficiency and applicability of Y-SSRs for genomic integration. In summary, this thesis made significant strides in the identification, characterization, and development of novel Y-SSRs for advanced genome engineering. The comprehensive prediction pipeline, combined with in-depth molecular characterization, has expanded the genetic toolbox to meet the growing demand for better genome editing tools. By exploring efficiency, cross-specificity, and potential cytotoxicity, this research lays the foundation for the safe and effective application of novel Y-SSRs in various therapeutic settings. Furthermore, by demonstrating the potential of these recombinases to improve efforts in creating designer recombinases through directed evolution, this research has opened new avenues for precise genome surgery. The successful development and implementation of these novel recombinases have the potential to revolutionize gene therapy, synthetic biology, and our understanding of gene function and regulation.

Cystic Fibrosis (CF) is a common life-shortening autosomal recessive disease that affects over 100.000 people worldwide people worldwide. It is caused by mutations in the CF trans-membrane conductance regulator (CFTR) gene, that encodes for a membrane channel localized at the apical surface of epithelial cells where it has a crucial role in the secretion of chloride and bicarbonate. Over 2100 different CFTR mutations have been reported and among the pathogenic once the most common is F508del, located in the nucleotide-binding domain 1 (NBD1). F508del is a three-nucleotide deletion that results in the loss of a phenylalanine at position 508 in the protein and in the consequent CFTR degradation by the ubiquitin-proteasome system. Different attempts to correct F508del-CFTR gene were made using genome editing approaches, however deletions like F508del remain difficult to be repaired. Several studies reported that additional mutations (revertant mutations) in the F508del-CFTR gene can rescue both CFTR folding and activity, suggesting a potential novel strategy to correct F508del. For this reason, the first aim of this work was the identification of novel F508del-CFTR revertants that can rescue CFTR localization and function. We generated a library of mutants introducing random substitutions into the F508del-CFTR gene. Revertant mutations were isolated based on their ability to rescue the presence of CFTR at the plasma membrane (PM) in HEK293T cells and identified by Sanger sequencing. Restoration of CFTR maturation, localization, and function of the identified revertants was evaluated by western blot, flow cytometry analysis and YFP assay, reaching levels similar to the wild type CFTR. Then we used CRISPR-Cas technology to introduce selected revertant mutations, such as I539T, R553Q, G550E, R555K and R1070W, in the endogenous F508del-CFTR gene. Adenine and cytosine base editors (ABE and CBE) allow the insertion of the desired base conversion without the formation of double strand breaks. Efficient editing was evaluated through Sanger sequencing, reaching up to 60% of base conversion. CFTR rescue at the PM in edited cells was analyzed by flow cytometry showing different degrees of recovery compared to the wild type CFTR. In this work, we confirmed that revertant mutations can rescue F508del CFTR localization and function. In addition, we demonstrated that CRISPR-base editors are valid tools to introduce these mutations in the endogenous F508del-CFTR gene, leading to a permanent correction. The proposed strategy could overcome the limits that genome editing strategies faced till now in the correction of F508del, providing a new potential therapeutic approach to treat CF.

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