Faculty Bibliography

Altered protein conformation can cause incurable neurodegenerative disorders. Mutations in SERPINI1, the gene encoding neuroserpin, can alter protein conformation resulting in cytotoxic aggregation leading to neuronal death. Familial encephalopathy with neuroserpin inclusion bodies (FENIB) is a rare autosomal dominant progressive myoclonic epilepsy that progresses to dementia and premature death. We developed HEK293T and induced pluripotent stem cell (iPSC) models of FENIB, harboring a patient-specific pathogenic SERPINI1 variant or stably overexpressing mutant neuroserpin fused to GFP (MUT NS-GFP). Here, we utilized a personalized adenine base editor (ABE)-mediated approach to correct the pathogenic variant efficiently and precisely to restore neuronal dendritic morphology. ABE-treated MUT NS-GFP cells demonstrated reduced inclusion size and number. Using an inducible MUT NS-GFP neuron system, we identified early prevention of toxic protein expression allowed aggregate clearance, while late prevention halted further aggregation. To address several challenges for clinical applications of gene correction, we developed a neuron-specific engineered virus-like particle to optimize neuronal ABE delivery, resulting in higher correction efficiency. Our findings provide a targeted strategy that may treat FENIB and potentially other neurodegenerative diseases due to altered protein conformation such as Alzheimer's and Huntington's diseases.

Unintended on-target chromosomal alterations induced by CRISPR/Cas9 in mammalian cells are common, particularly large deletions and chromosomal translocations, and present a safety challenge for genome editing. Thus, there is still an unmet need to develop safer and more efficient editing tools. We screened diverse DNA polymerases of distinct origins and identified a T4 DNA polymerase derived from phage T4 that strongly prevents undesired on-target damage while increasing the proportion of precise 1- to 2-base-pair insertions generated during CRISPR/Cas9 editing (termed CasPlus). CasPlus induced substantially fewer on-target large deletions while increasing the efficiency of correcting common frameshift mutations in DMD and restored higher level of dystrophin expression than Cas9-alone in human cardiomyocytes. Moreover, CasPlus greatly reduced the frequency of on-target large deletions during mouse germline editing. In multiplexed guide RNAs mediating gene editing, CasPlus repressed chromosomal translocations while maintaining gene disruption efficiency that was higher or comparable to Cas9 in primary human T cells. Therefore, CasPlus offers a safer and more efficient gene editing strategy to treat pathogenic variants or to introduce genetic modifications in human applications.

To solve the clinical transformation dilemma of lamin A/C (LMNA)-mutated dilated cardiomyopathy (LMD), we developed an LMNA-mutated primate model based on the similarity between the phenotype of primates and humans. We screened out patients with LMD and compared the clinical data of LMD with TTN-mutated and mutation-free dilated cardiomyopathy to obtain the unique phenotype. After establishment of the LMNA c.357-2A>G primate model, primates were continuously observed for 48 months, and echocardiographic, electrophysiological, histologic, and transcriptional data were recorded. The LMD primate model was found to highly simulate the phenotype of clinical LMD. In addition, the LMD primate model shared a similar natural history with humans.

Mutations of PTEN-induced kinase I (PINK1) cause early-onset Parkinson's disease (PD) with selective neurodegeneration in humans. However, current PINK1 knockout mouse and pig models are unable to recapitulate the typical neurodegenerative phenotypes observed in PD patients. This suggests that generating PINK1 disease models in non-human primates (NHPs) that are close to humans is essential to investigate the unique function of PINK1 in primate brains. Paired single guide RNA (sgRNA)/Cas9-D10A nickases and truncated sgRNA/Cas9, both of which can reduce off-target effects without compromising on-target editing, are two optimized strategies in the CRISPR/Cas9 system for establishing disease animal models. Here, we combined the two strategies and injected Cas9-D10A mRNA and two truncated sgRNAs into one-cell-stage cynomolgus zygotes to target the PINK1 gene. We achieved precise and efficient gene editing of the target site in three newborn cynomolgus monkeys. The frame shift mutations of PINK1 in mutant fibroblasts led to a reduction in mRNA. However, western blotting and immunofluorescence staining confirmed the PINK1 protein levels were comparable to that in wild-type fibroblasts. We further reprogramed mutant fibroblasts into induced pluripotent stem cells (iPSCs), which showed similar ability to differentiate into dopamine (DA) neurons. Taken together, our results showed that co-injection of Cas9-D10A nickase mRNA and sgRNA into one-cell-stage cynomolgus embryos enabled the generation of human disease models in NHPs and target editing by pair truncated sgRNA/Cas9-D10A in PINK1 gene exon 2 did not impact protein expression.

There are an estimated 10 000 monogenic diseases affecting tens of millions of individuals worldwide. The application of CRISPR/Cas genome editing tools to treat monogenic diseases is an emerging strategy with the potential to generate personalized treatment approaches for these patients. CRISPR/Cas-based systems are programmable and sequence-specific genome editing tools with the capacity to generate base pair resolution manipulations to DNA or RNA. The complexity of genomic insults resulting in heritable disease requires patient-specific genome editing strategies with consideration of DNA repair pathways, and CRISPR/Cas systems of different types, species, and those with additional enzymatic capacity and/or delivery methods. In this review we aim to discuss broad and multifaceted therapeutic applications of CRISPR/Cas gene editing systems including in harnessing of homology directed repair, non-homologous end joining, microhomology-mediated end joining, and base editing to permanently correct diverse monogenic diseases.

Many human genetic diseases, including Hutchinson-Gilford progeria syndrome (HGPS), are caused by single point mutations. HGPS is a rare disorder that causes premature aging and is usually caused by a de novo point mutation in the LMNA gene. Base editors (BEs) composed of a cytidine deaminase fused to CRISPR/Cas9 nickase are highly efficient at inducing C to T base conversions in a programmable manner and can be used to generate animal disease models with single amino-acid substitutions. Here, we generated the first HGPS monkey model by delivering a BE mRNA and guide RNA (gRNA) targeting the LMNA gene via microinjection into monkey zygotes. Five out of six newborn monkeys carried the mutation specifically at the target site. HGPS monkeys expressed the toxic form of lamin A, progerin, and recapitulated the typical HGPS phenotypes including growth retardation, bone alterations, and vascular abnormalities. Thus, this monkey model genetically and clinically mimics HGPS in humans, demonstrating that the BE system can efficiently and accurately generate patient-specific disease models in non-human primates.

Duchenne muscular dystrophy (DMD) is a fatal genetic disorder caused by mutations in the dystrophin gene. To enable the non-invasive analysis of DMD gene correction strategies in vivo, we introduced a luciferase reporter in-frame with the C-terminus of the dystrophin gene in mice. Expression of this reporter mimics endogenous dystrophin expression and DMD mutations that disrupt the dystrophin open reading frame extinguish luciferase expression. We evaluated the correction of the dystrophin reading frame coupled to luciferase in mice lacking exon 50, a common mutational hotspot, after delivery of CRISPR/Cas9 gene editing machinery with adeno-associated virus. Bioluminescence monitoring revealed efficient and rapid restoration of dystrophin protein expression in affected skeletal muscles and the heart. Our results provide a sensitive non-invasive means of monitoring dystrophin correction in mouse models of DMD and offer a platform for testing different strategies for amelioration of DMD pathogenesis.

Muscular dystrophies represent a large group of genetic disorders that significantly impair quality of life and often progress to premature death. There is no effective treatment for these debilitating diseases. Most therapies, developed to date, focus on alleviating the symptoms or targeting the secondary effects, while the underlying gene mutation is still present in the human genome. The discovery and application of programmable nucleases for site-specific DNA double-stranded breaks provides a powerful tool for precise genome engineering. In particular, the CRISPR/Cas system has revolutionized the genome editing field and is providing a new path for disease treatment by targeting the disease-causing genetic mutations. In this review, we provide a historical overview of genome-editing technologies, summarize the most recent advances, and discuss potential strategies and challenges for permanently correcting genetic mutations that cause muscular dystrophies.

Genome editing with CRISPR/Cas9 is a promising new approach for correcting or mitigating disease-causing mutations. Duchenne muscular dystrophy (DMD) is associated with lethal degeneration of cardiac and skeletal muscle caused by more than 3000 different mutations in the X-linked dystrophin gene (DMD). Most of these mutations are clustered in "hotspots." There is a fortuitous correspondence between the eukaryotic splice acceptor and splice donor sequences and the protospacer adjacent motif sequences that govern prokaryotic CRISPR/Cas9 target gene recognition and cleavage. Taking advantage of this correspondence, we screened for optimal guide RNAs capable of introducing insertion/deletion (indel) mutations by nonhomologous end joining that abolish conserved RNA splice sites in 12 exons that potentially allow skipping of the most common mutant or out-of-frame DMD exons within or nearby mutational hotspots. We refer to the correction of DMD mutations by exon skipping as myoediting. In proof-of-concept studies, we performed myoediting in representative induced pluripotent stem cells from multiple patients with large deletions, point mutations, or duplications within the DMD gene and efficiently restored dystrophin protein expression in derivative cardiomyocytes. In three-dimensional engineered heart muscle (EHM), myoediting of DMD mutations restored dystrophin expression and the corresponding mechanical force of contraction. Correcting only a subset of cardiomyocytes (30 to 50%) was sufficient to rescue the mutant EHM phenotype to near-normal control levels. We conclude that abolishing conserved RNA splicing acceptor/donor sites and directing the splicing machinery to skip mutant or out-of-frame exons through myoediting allow correction of the cardiac abnormalities associated with DMD by eliminating the underlying genetic basis of the disease.