Browsing by Subject "Genome editing"

Successful transfection of genetically active materials is essential to gene delivery and genome editing. Electrotransfection, also known as electroporation, is a fast, safe, and convenient non-viral method for introducing materials such as proteins and nucleic acids into cells and tissues. It has been widely used in academic research, industrial manufacturing, and clinical therapeutics. Particularly, electrotransfection is one of the most commonly used method in gene delivery into mammalian cells. However, despite its many advantages comparing to other gene delivery methods, the application of electrotransfection is limited by inconsistent transfection efficiency, which is caused by the poor understanding of the mechanism of electrotransfection.

The goal of my research is to understand the fundamental biological mechanisms of electrotransfection and to develop novel strategies that can improve the transfection efficiency of gene delivery and genome editing. To this end, this study is divided into two phases. Phase 1 aims at understanding the key cellular components involved in the transport process. Phase 2 focuses on the development of strategies to enhance electrotransfection by controlling the biological pathways that are involved in electrotransfection.

In the first phase of my study, we investigated the dependence of electrotransfection efficiency on endocytosis. Data from this study demonstrated that macropinocytosis is involved in electrotransfection. The results revealed that electric pulses induced cell membrane ruffling and actin cytoskeleton remodeling. Using fluorescently labeled pDNA and a macropinocytosis marker (i.e., dextran), the study showed that electrotransfected pDNA co-localized with dextran in intracellular vesicles formed from macropinocytosis. Furthermore, electrotransfection efficiency was reduced significantly by lowering temperature or treatment of cells with a pharmacological inhibitor of Rac1 and could be altered by changing Rac1 activity. Taken together, the findings suggested that electrotransfection of pDNA involved Rac1-dependent macropinocytosis.

Second phase of this study focuses on the intracellular transport of plasmid DNA, especially the transport of DNA molecules towards degradative compartments. Our data elucidated that components in both endocytic and autophagic pathways are responsible for intracellular trafficking and processing of transfected materials such as pDNA. In addition, we also characterized a new type of vesicle named amphisome-like vesicle (ALB) and revealed its involvement in electrotransfection. Based on these findings, we propose a novel strategy to enhance electrotransfection by blocking degradative routes within the endocytic pathways, which led to the development of a new technique called transfection by redirection of endocytic and autophagic traffic (TREAT). Transfection of plasmid DNA (pDNA), messenger RNA (mRNA), sleeping beauty transposon system (SB), and different forms of CRISPR/Cas9 system by TREAT achieved superior efficiency in various cell lines including difficult-to-transfect human primary cells. In addition, we successfully applied TREAT method to improve clinically relevant applications including SB-based gene integration and CRISPR/Cas9-based editing of T cell receptor alpha constant (TRAC). In summary, we studied the biological mechanism of electrotransfection and developed a general, flexible, and reliable technique to enable highly efficient non-viral gene delivery and genome editing. Furthermore, the insights gained on the mechanism of electrotransfection provide better understanding of cellular response to exogenous materials. In the future, our study could potentially pave new paths for a wide range of research and therapeutic applications such as CRISPR/Cas9 mediated high-throughput loss-of-function gene screening analysis, correction of disease-related mutations, as well as genetic engineering of immune cells and stem cells for transplantation.

Therapeutic gene transfer and genome editing require effective delivery of genetic cargo to target cells and tissues. Recombinant adeno-associated viral (AAV) vectors are a promising delivery platform, but ongoing clinical trials continue to highlight a relatively narrow therapeutic window. Efforts to optimize vector dosing or engineer improved vectors are confounded, at least in part, by differences in AAV biology across animal species. Here, we present a broadly applicable, cross-species evolution approach to tackle this challenge. Specifically, I iteratively cycled AAV libraries administered intravenously and amplified isolates from CNS tissue in pigs, mice, and non-human primates to generate cross-species compatible AAVs (ccAAVs). By sequentially evolving AAV libraries in three different species, we discover a highly potent variant (AAV.cc47) that demonstrates improved attributes benchmarked against AAV serotype 9 (AAV9). Increased potency of AAV.cc47 is evidenced through robust reporter gene expression as well as Cre-mediated recombination and CRISPR/Cas9-mediated genome editing in a fluorescent reporter mouse model. Enhanced transduction efficiency of AAV.cc47 vectors is further corroborated in macaques and pigs, providing a strong rationale for potential clinical translation into human gene therapies. Lastly, we report increased expression of a therapeutic acid alpha-glucosidase (GAA) transgene in a mouse model of Pompe disease and enhanced restoration of dystrophin through CRISPR/Cas9 gene editing in the mdx mouse model of Duchenne Muscular Dystrophy using AAV.cc47 vectors. We discovered another cross-species compatible variant (AAV.cc84) with enhanced CNS transduction and de-targeted from the liver in vivo compared to AAV9. We demonstrate improved targeting of neurons in the brain following both systemic and intracerebroventricular (ICV) injection of reporter vectors in mice. Reporter gene expression and vector genome biodistribution reveal a liver detargeted phenotype with AAV.cc84 compared to AAV9. Lastly, enhanced neuronal transduction was confirmed in the pig CNS following intrathecal infusion of AAV.cc84 reporter vectors. Taken together, we envision that ccAAV vectors can potentially improve predictive modeling in preclinical studies as well as clinical translatability by broadening the therapeutic window of AAV based gene therapies.

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.

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.

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.

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.

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.

Glycogen storage disease type Ia is an autosomal recessive disorder caused by a mutation in the glucose-6-phosphatase (G6Pase) catalytic subunit, encoded in humans by G6PC. G6Pase dephosphorylates glucose-6-phosphate (G6P) in the liver to generate glucose that can be shuttled to the bloodstream to maintain normoglycemia. Patients with GSD Ia typically present at 6 months of age with sever hypoglycemia, which is lethal if untreated. The current treatment is a strict dietary regimen in which children must be fed every 2 hours overnight or given nasogastric tube feeding, and adults must consume uncooked cornstarch around the clock to maintain normal blood sugar levels. This treatment maintains survival but fails to prevent other symptoms related to metabolism of the excess G6P, and patients develop hepatic adenomas that may become hepatocellular carcinoma later in life, in addition to progressive renal complications.

To overcome the problems persisting during dietary therapy, the Koeberl lab has sought to develop gene therapy approaches that use adeno-associated virus (AAV) vectors to replace the G6pase activity, restoring normoglycemia and normal metabolic processes. However, the vast majority of AAV-delivered genetic material exists as episomes that do not replicate as cells divide, so the effects of AAV gene therapy on GSD Ia mouse and dog models have proven temporary. We hypothesized that driving integration of therapeutic vector genomes into an affected individual's genome would improve beneficial effects' longevity.

We tested several approaches to accomplish this, and have found positive effects using a zinc finger nuclease (ZFN) that targets the mouse safe harbor ROSA26 locus to induce homologous recombination of the G6PC donor vector into the mouse genome. We were able to see an improvement in mouse survival to 8 months of age, an increase in G6Pase activity at 3 months of age, and a decrease in glycogen accumulation at 3 months of age, when the ZFN vector is administered alongside the G6PC vector, compared with mice that received the G6PC vector alone.

We have also taken an alternative approach to overcoming the long-term complications of the current dietary treatment, which would augment rather than replace the current treatment. We have examined several drugs known to induce autophagy in other disease models or cell culture systems, to determine if we could manipulate autophagic activity in G6PC knockdown hepatocytes or GSD Ia mice. We have found positive results using rapamycin, a well-studied MTOR inhibitor, in mice and cells, and have screened several other drugs as well, finding positive effects for bezafibrate, mifepristone, carbamazepin, and lithium chloride, in terms of lipid reduction (which accumulates as a symptom of GSD Ia) and/or LC3-II enhancement, which is reduced in GSD Ia due to downregulation of autophagy during G6P accumulation.

Glycogen storage disease type Ia (GSD Ia) is an autosomal recessive metabolicdisorder caused deficiency of glucose-6-phosphatase (G6Pase) resulting from pathogenic variants in the G6PC gene. G6Pase catalyzes the hydrolysis of glucose-6-phosphate to release glucose which can then enter the bloodstream. GSD Ia patients have excess glycogen accumulation mainly in the liver and kidneys and suffer from life threatening hypoglycemia. The current treatment for GSD Ia is dietary therapy that requires patients to frequently consume uncooked cornstarch on a strict schedule. Cornstarch provides a complex carbohydrate that slowly releases glucose to prevent hypoglycemia. This treatment fails to prevent long-term complications associated with GSD Ia including renal failure and the development of hepatocellular adenomas and carcinomas. This lab and others have developed adeno-associated virus (AAV) vector based gene therapies to deliver and therapeutic G6PC transgene to affected tissues in GSD Ia animal models. However, the therapeutic effect is limited as AAV vector genomes are rapidly lost and the biochemical correction declines. Currently no treatment for GSD Ia exists that provides stable, robust expression of G6Pase that can clear glycogen and prevent hypoglycemia. This study employed a novel genome editing approach designed to insert the therapeutic G6PC into the endogenous locus in canine and murine models of GSD Ia. Integration of the transgene into the genome will promote stable expression of G6Pase and prevent the decline of vector genomes and the therapeutic benefit. This genome editing approach utilizes the CRISPR/Cas9 system to generated targeted double stranded DNA breaks at a targeted site in the genome. The G6PC transgene is present in a Donor template with homology to the DNA break to drive homology directed repair (HDR) resulting in the integration of the transgene into the genome. In a canine model of GSD Ia, editing and incorporation of the transgene was achieved in both adult dogs and puppies. Up to 1.0% of alleles were edited in the dog livers and contained the transgene. G6Pase production from the integrated transgene was detected, which correlated with prevention of hypoglycemia during fasting. This demonstrated genome editing in the liver of a large animal model for an inherited metabolic disorder using HDR to insert a therapeutic transgene. A subsequent study in GSD Ia mice also showed incorporation of a G6PC transgene in the mouse genome. Mice were treated with either the Donor transgene vector alone or with both the Donor and a CRISPR/Cas9 vector to assess to role of nuclease activity on integration. Mice treated with both vectors demonstrated improved blood glucose concentrations during fasting, decreased liver glycogen, and increased vector genome copies. Treatment with the pan PPAR agonist bezafibrate increased the efficiency of genome editing. Mice treated with bezafibrate that received both editing vectors had 5.9% of alleles that contained the integrated transgene, whereas only 3.1% of alleles contained the transgene in mice not treated with the drug. This work showed that integration of a therapeutic transgene using CRISPR/Cas9 based genome editing is possible in murine and canine models of GSD Ia. Editing resulted in biochemical correction and sustained transgene expression. These data support the further development of genome editing technologies for GSD Ia and other inherited metabolic disorders.