DNA sequence information is constantly threatened by damage. In the clinic, intentional DNA damage is often used to treat cancer. Cisplatin, a first-line chemotherapy used to treat millions of patients, functions specifically by generating physical links within DNA strands, blocking DNA replication, and killing dividing cells. To maintain genome integrity, organisms have evolved the capacity to repair, respond, or otherwise resist change to the DNA sequence through a network of genetically encoded DNA damage tolerance pathways. In chapter 1, I present advances in experimental design and current progress for a systems genetics approach, using Saccharomyces cerevisiae, to reveal relationships between cisplatin tolerance pathways. Additionally, recent efforts to sequence thousands of cancer genomes have revealed recurrent genetic changes that cause overexpression of specific cisplatin tolerance genes. In chapter 2, I present a submitted manuscript that models overexpression of an essential cisplatin tolerance gene. This study uses a systems genetics approach to reveal the genetic pathways that are essential for tolerating this perturbation, which ultimately led to mechanistic insights for this gene. Convenient genome engineering in Saccharomyces has made this organism an ideal model to develop systems genetics concepts and approaches. In chapter 3, I present a published manuscript that demonstrates a new approach to disrupting genes by making site-specific nonsense mutations. Importantly, this approach does not require cytotoxic double-strand DNA breaks and is applicable to many model organisms for disrupting almost any gene, which may advance systems genetics into new model organisms. Systems genetics provides a framework for determining how DNA damage tolerance pathways act together to maintain cellular fitness and genome integrity. Such insights may one day help clinicians predict which cancers will respond to treatment, potentially sparing patients from unnecessary chemotherapy.

The question of delivery has become a critical part of therapeutic research and development. While nanoparticle formulations are now used in a variety of FDA-approved therapies, these focus on simple oral formulations or systemic lipid nanoparticle administration; therefore, significant research is ongoing in the development of specialized delivery systems for more complex cargos or specific targeting[1]. For therapeutics that have high potential for toxicity, like chemotherapeutics, delivery systems need to transport disproportionately into target organs and further target cells. This is more so true for therapeutics that require delivery to a specific cellular compartment to have significant efficacy, like in the case of gene editing in the nucleus or mitochondria-targeted peptides. To contribute to the development of efficient carriers for gene and drug therapy, this work aims to explore the potential of liposome-coated mesoporous silica nanoparticles and engineering methods that provide functionality to defeat barriers to efficient therapeutic development. Recent advances in CRISPR/Cas9 technology present an attractive toolset to study genes for the development of novel therapeutics. Since traditional delivery methods for gene therapies relied heavily on viruses— requiring biosafety level clearance and eliciting immunogenicity concerns, as well as limitations in multiplex gene editing capabilities in single viral vehicles— recently, nanoparticles have become an attractive nonviral alternative for gene therapy research and development. While lipid-based nanoparticles have been at the forefront of siRNA and mRNA therapies, we looked at increasing the loading capabilitiesof a liposome by using a mesoporous silica nanoparticle (MSN) core. The MSN provides efficient electrostatic loading of the relatively large and non-uniformly charged CRISPR/Cas9 protein and guide RNA ribonucleoprotein complex (RNP) as well as more charge-dense plasmids, while a liposome coating offers the PEGylation and targeting capabilities necessary for selective uptake and in vivo application. After demonstrating gene-editing efficiencies above 20% for both plasmid and RNP modalities of CRISPR/Cas9, we tested its application in multiplex gene editing for the study lipid metabolism pathways. To demonstrate the maintained efficiency of this system, this liposome-coated MSN (lipoMSN) platform was used to deliver a combination of RNPs targeting three genes involved in lipid metabolism in the liver. These genes, Pcsk9, Apoc3, and Angptl3, are derived from research demonstrating that populations with loss-of-function mutations in any one of these genes garner improved cardiovascular health, characterized by lowered blood cholesterol and triglycerides. The lipoMSN system demonstrated that it maintained significant gene editing, above 25% efficiency, at a specific gene target despite reduced dosage of target-specific RNP due to the combination of other target RNP. By leveraging this system to deliver various combinations of targeting RNPs in the same nanoparticle and therefore ensuring a higher probability that any given cell is edited at all targets, synergistic effects on lipid metabolism can be observed in vitro and in vivo. These effects, such as an approximately 50% decrease in serum LDL-cholesterol 4-weeks after treatment with pcsk9 and angptl3- targeted RNPs, have not been observed in previous studies. Continued work with this lipoMSN platform is ongoing, with projects leveraging the system for multiplex gene disruption as well as endosomal delivery of peptide and chemical drugs. We are currently leveraging the comparatively low spread of lipoMSN to provide gene disruption of adra1a, adra1b, and adra1d in the discrete area of the thalamus. Further, this system is also well suited for other CRISPR/Cas9 elements, such as deactivated Cas9 (dCas9) fused to epigenetic modifiers, enhancers and repressors. To demonstrate another facet of the lipoMSN delivery platform, we adjusted the formulation for the specific delivery to endosomal compartments of nociceptive neurons. Previous work by our collaborators at the Bunnett Lab provided evidence that signaling in the endosomal compartment is partially responsible for both pain propagation and amelioration. In order to provide highly targeted treatment to reduce the off-target effects linked with opioid-based pain treatment, we wanted to leverage the resonance of these nanoparticles in the endosomes following endocytosis and enhance the drug delivery to the endosomal compartments propagating nociceptive signaling. We did this by first integrating a targeting ligand in the form of a DADLE-peptide, to the outside of the liposomes, providing a 20-40% increase in uptake for delta-opioid receptor (DOR)-expressing cell types. Further, we used an oxidation- and pH-sensitive MSN core to provide increased drug release to the endosomal environment, which is naturally oxidizing and at a lower pH of approximately 5.2. This resulted in an increased therapeutic effect when compared to naked peptide drug in a mouse model of neurogenic pain. We are also looking at leveraging this system for other endosomal signaling pathways and applying our lipoMSN platform to cargos such as glucagon-like peptide-1 (GLP-1) receptor agonists for diabetes and US28 receptor antagonists for reduction of proliferative signaling in cancers. Collectively, these projects provide insight on how to design delivery vehicles for specific gene and drug delivery. This lipoMSN system has the potential to be a versatile platform for the development of combinatorial gene therapeutics in liver-related disease. Further, this platform may inform research and development in the next generation of endosomally-targeted therapeutics for increased efficacy and reduced off-target or side effects.

Rare gene-disrupting variants and common regulatory variants play key roles in rare and common disease, respectively. These variants are of great interest for investigation into genetic contributions to disease, but experimental methods to validate their impact on gene expression levels are lacking. In this study, we utilized CRISPR/Cas9 genome editing to validate regulatory variants including cis-eQTLs, rare stop-gained variants in healthy and disease cases and one immune-response trans-eQTL master regulator. For investigation into common and rare regulatory variants within transcribed regions, we developed a scalable CRISPR-based polyclonal assay for experimental assessment. First, we applied this assay to nine rare stop-gained variants found in the general population, in GTEx. After editing, the stop-gained variants show a significant allele-specific depletion in transcript abundance, as expected. Next, we utilized the assay to validate 33 common eQTLs found in GTEx. After editing, the eQTL variants show higher variance in effect size than control variants, indicating a regulatory effect. Finally, we applied the polyclonal editing approach to clinical and new stop-gained variants in two disease-associated genes. The results follow the expected trend, with NMD being triggered by variants upstream of the NMD threshold but not by those beyond. This method demonstrates scalable experimental confirmation of putative causal regulatory variants, and improved interpretation of regulatory variation in humans. Next, we sought to experimentally validate an immune-response eQTL for IRF1 in cis and many genes in trans under LPS stimulation. We used CRISPRi to repress the enhancer locus and found that the enhancer is active in our immune cell system. Next, we used CRISPR-Cas9 genome editing and isolation of monoclonal cell lines to target this variant locus. After LPS stimulation, we performed RNA-sequencing on wild type and edited clones, showing that the effect size of the genes which are associated with the trans-eQTL are correlated with differential expression between the edited and wild type cell lines for the same genes. Additionally, we find that the differential expression between edited clones is correlated with CRISPRi repression of the IRF1 promoter and enhancer. In this way, we were able to identify a common genetic variant which modifies the transcriptomic immune response to LPS and validate the trans-eQTL signal.

Bacterial species and communities play foundational roles in human health and therapeutics, in vital ecological and environmental processes, and in industrial applications for the biosynthesis of valuable compounds and materials. However, existing genetic engineering methods and technologies available for bacterial functional genetics or large-scale genomic integration are inefficient, unable to translate between different target species, or lacking precise targeting or reprogramming capabilities. In this work, we describe a novel class of CRISPR- associated transposons (CRISPR-Tn) that facilitate programmable RNA-guided DNA insertions. In particular, the Tn6677 CRISPR-Tn system from Vibrio cholerae comprises a Tn7-like transposase machinery that has co-opted a nuclease-deficient Type I-F3 CRISPR-Cas system to guide its target selection. We show that, similar to canonical CRISPR-Cas systems, this CRISPR- Tn system can be easily programmed using the CRISPR RNA (crRNA) spacer sequence, and directs highly target-specific DNA integration into the Escherichia coli genome. After defining their core biological and mechanistic principles, we developed these CRISPR-Tn systems into a genome engineering platform, which we named INTEGRATE (Insertions of Transposable Elements by Guide RNA-Assisted Targeting). Particularly, optimization of V. cholerae Tn6677 (Vch INTEGRATE, or VchINT) produced a system capable of programmable, broad-bacterial- host, and multiplexed integration of DNA payloads up to 10 kilobases in length, with genomic editing efficiencies reaching 100%. Our single-plasmid expression of system components enabled, for the first time, genome engineering of specific target strains within a complex fecal bacterial community. In addition, we performed extensive deep sequencing within transposition experiments to characterize and examine non-conventional transposition products, including cointegrates formed through replicative transposition, and long-range integration events resulting from on-target DNA binding. Finally, by individually inserting transposon ends into the E. coli genome, we demonstrated successful transposition-mediated mobilization of a genomic fragment 100 kilobases (kb) in length, demonstrating engineering at the genome-scale using VchINT. Altogether, this work highlights the potential of VchINT and other CRISPR-Tn systems as next- generation genome engineering technologies in bacteria and beyond.

The world of precision medicine was revolutionized by the discovery of CRISPR-Cas systems. In particular, the capabilities of the programmable nuclease Cas9 and its derivatives have unlocked a world in which applied genome engineering to cure human disease is a reality being pursued in patient clinical trials. Gene editing via the induction of programmable, site-specific double strand breaks (DSBs) has been revolutionary for the precision medicine field. However, there are many safety concerns centered on the induction of DSBs causing potential undesirable on- and off-target consequences, particularly for in vivo CRISPR applications. To circumvent these warranted concerns, many groups have attempted to repurpose recombinases or engineer new fusion systems to perform programmable genome engineering without the induction of DSBs. This dissertation will first highlight the development of recombinases for programmable DNA insertions over the course of decades, including efforts to evolve novel DNA recognition sequences, efforts to tether recombinases to programmable DNA-binding proteins, and the recent discovery of naturally occurring RNA-guided DNA transposition systems. This dissertation will then highlight the development of CRISPR-associated transposases (CASTs) as DSB-independent programmable mammalian gene editing tools capable of integrating large DNA cargos, as well as the future directions that may further enhance CAST activity in human cells. The works in this dissertation detail the initial efforts to engineer and optimize a new class of genome manipulation tools that were previously absent from the gene editing toolkit.