Browsing by Subject "CRISPR-Cas9"

Spinal muscular atrophy of Jokela type (SMAJ) is an autosomal dominant motor-neuron disease caused by a missense mutation c.197G>T, p.G66V in the gene CHCHD10. Coiled-coil-helix-coiled-coil-helix domain-containing protein 10 (CHCHD10) is a nuclear-encoded mitochondrial protein located in the intermembrane space (IMS) of mitochondria with an unknown exact function and disease-causing mechanism. In this project, the overarching aim was to correct a heterozygous SMAJ-causing mutation in patient myoblast cells with CRISPR-Cas9 genome editing. The goal was to create a genetically identical, isogenic, cell line to study only the effects of the mutation on cellular phenotype in vitro. Human myoblast cells isolated from patient biopsies provide the most pertinent experimental model to study neuromuscular atrophy-associated mutations in their natural genomic environment. More specific aims included genome editing optimization with myoblast cells, since it is not as widely conducted as with some other cell types, such as iPSCs. CRISPR-Cas9 ribonucleoprotein (RNP) complex and associated donor template were used to induce homology-directed repair (HDR) in the genome of patient-derived myoblast cells and correct the mutation. After optimization of electroporation conditions for myoblast cells, guide RNAs were designed and transfected into patient myoblasts. Clonal cell lines were made by utilizing techniques such as fluorescence adjusted cell sorting (FACS) and manual colony picking. The success and precision of genome editing were analyzed by Sanger sequencing, comparing the performance of the different guide RNAs with restriction enzyme analysis and Synthego ICE CRISPR web tool, and screening regions of potential off-target genome editing. A genome-edited myoblast cell line with the CHCHD10 c.197G>T mutation corrected, was successfully generated to provide an isogenic control for the patient myoblast cell line. Optimization of myoblast electroporation was successful and conditions used proved to be effective. Clonal cell line creation proved to be challenging with myoblast cells and work is still needed to improve the viability of single-cell clones after FACS. Nevertheless, the advances taken here regarding myoblast genome editing with CRISPR-Cas9 offer a fertile avenue for future research of myoblasts genome manipulation, myogenic disorders, and the role of CHCHD10 in skeletal muscle and SMAJ. Comparing the CHCHD10 protein level and mRNA expression between patient cells, corrected myoblasts, and differentiated myotubes is an area of future research. Future work also includes measuring the mitochondrial integrated stress response in both cell lines and co-culturing myotubes and iPSC derived motor neurons to study the effects of p.G66V on neuromuscular junction (NMJ) formation.

As a genome editing tool, CRISPR-Cas9 has provided a robust way to generate mutations in the gene of interest, at a certain time point, and in selected cell populations. The impairment of dopaminergic neurons in the substantia nigra is addressed to be one of the main pathologies of Parkinson’s disease. The histopathology of Lewy Bodies, with an undetermined role, accompanies the demise of DA neurons. Development of strategies for the prevention the neurodegeneration has a potential to slow down the progression of Parkinson’s disease. In this study, a novel, neuron-specific CRISPR-Cas9 system was developed for the purpose of dissecting neuroprotective pathways in primary dopaminergic neurons. The optimization of the tool was done by targeting EGFP at TH-positive neurons obtained from transgenic animals expressing EGFP in dopaminergic neurons. Complete loss of EGFP was achieved at day 6 after the introduction of the CRISPR-Cas9 via lentiviral vectors. There were no survival or transduction efficiency differences. Two significant pathways for the survival of dopaminergic neurons, the microRNA biogenesis and GDNF/RET signaling were selected to collect the preliminary data. Dicer, Trbp, Translin, Ago-2 and Ret were targeted with single sgRNAs, which were specifically designed to create indel mutations in these genes, and specific lentivirus vectors were produced with each guide. After transduction with the lentivirus vectors, survival of the TH-positive neurons was unaffected. Data obtained from the quantitative PCR suggested that there was 50-70% decline in transcript levels of Trbp. However, the unchanged transcript levels of the other miRNA-related targets suggest the need for further optimization of the specific guides. Knockdown of Ret was validated by inhibition of pharmacological benefits of GDNF. Overall, this research has shown the further development of this CRISPR-Cas9 tool would be useful to dissect neuroprotective signaling pathways in dopaminergic neurons.

The Clustered Regularly-Interspaced Short Palindromic Repeats (CRISPR) and CRISPR associated protein (Cas9) (CRISPR-Cas9) system is a widely used gene editing technology due to its potential to alter the genome precisely in desired locations. Due to the potential of the CRISPR-Cas9 system, the objective of the thesis is to improve the precise editing of genes by modifying the CRISPR-Cas9 platform. Ultimately, the aim is to develop a platform that can edit any mutation and repair it to a normal, functional gene in patient cells. In general, CRISPR-Cas9 provides opportunities in treating monogenic diseases, for example by modifying long-term hematopoietic stem cells in immunodeficiencies. CRISPR-Cas9 can target disease-causing mutation sites and introduce double-strand breaks. Afterwards, the native DNA repair machinery of a cell repairs the cut site either by more efficient, error-prone non-homologous end joining (NHEJ) or precise homology-directed recombination (HDR). In most clinically oriented genome editing studies, the desired repair outcome is the latter because it allows precise repair of the mutation according to the exogenous repair template. Despite all its positive features, the optimization of CRISPR-based editing system is crucial before medical use; CRISPR-Cas9 induces a p53-mediated DNA damage response, which leads to a transient G1 cell cycle arrest and hampers HDR-based precision genome editing. Other problems include the repair pathway depending on the cell cycle phase, repair template proximity, and off-target activity. This thesis demonstrates that Cas9 fusions allow addressing the problems mentioned above. Cas9 fusions with DNA repair proteins ensure improved editing efficiency at the close proximity to the target site in HEK293T, BJ5-ta and RPE reporter cell lines. In addition, Cas9 coupled with the engineered cell cycle timer, AcrⅡA2-cdt1, favors the editing at the S/G2 cell cycle phases avoiding the p53-mediated response. AcrⅡA2-cdt1 is a reversible, phage-derived CRISPR inhibitor that selectively inhibit CRISPR-Cas9 at the G1 cell cycle phase and releasing it at the S phase. This thesis provides extensive look on the CRISPR-Cas9 editing and its challenges in immortalized cell lines and primary cells. In the thesis, the generation of reporter cell lines is prior to the validation of the novel Cas9-fusions. Furthermore, the optimization of primary T cell and CD34+ hematopoietic stem cell electroporation with different electroporation systems brings the study closer to clinical applications. The thesis provides insights about the effect of the target site and the cell type for genome editing outcomes. The editing efficiencies depend on the Cas9 fusion protein, cell type and its proliferation rate. The editing efficiency in primary T cells and CD34+ hematopoietic stem cells can significantly improve by optimizing transfection and culturing conditions, such as concentration of the CRISPR-Cas9 complex, cell culturing time and electroporation program. Cas9 fusions improve the safety and efficiency of the CRISPR-Cas9 system depending the cell type and the proliferation rate of the cell. Timing the induction of double-strand breaks also improves the editing efficiency. Overall, the methods used in the thesis give useful tools for eventual translational applications.