Academic literature on the topic 'CRISPR-Cas9, genome editing, CDKL deficiency disorder'

Ornithine transcarbamylase (OTC) deficiency is an X-linked urea cycle disorder associated with high mortality. Although a promising treatment for late-onset OTC deficiency, adeno-associated virus (AAV) neonatal gene therapy would only provide short-term therapeutic effects as the non-integrated genome gets lost during hepatocyte proliferation. CRISPR-Cas9-mediated homology-directed repair can correct a G-to-A mutation in 10% of OTC alleles in the livers of newborn OTC spfash mice. However, an editing vector able to correct one mutation would not be applicable for patients carrying different OTC mutations, plus expression would not be fast enough to treat a hyperammonemia crisis. Here, we describe a dual-AAV vector system that accomplishes rapid short-term expression from a non-integrated minigene and long-term expression from the site-specific integration of this minigene without any selective growth advantage for OTC-positive cells in newborns. This CRISPR-Cas9 gene-targeting approach may be applicable to all patients with OTC deficiency, irrespective of mutation and/or clinical state.

Fabry nephropathy is a major cause of morbidity and premature death in patients with Fabry disease (FD), a rare X-linked lysosomal storage disorder. Gb3, the main substrate of α-galactosidase A (α-Gal A), progressively accumulates within cells in a variety of tissues. Establishment of cell models has been useful as a tool for testing hypotheses of disease pathogenesis. We applied CRISPR/Cas9 genome editing technology to the GLA gene to develop human kidney cell models of FD in human immortalized podocytes, which are the main affected renal cell type. Our podocytes lack detectable α-Gal A activity and have increased levels of Gb3. To explore different pathways that could have distinct patterns of activation under conditions of α-gal A deficiency, we used a high-throughput antibody array to perform phosphorylation profiling of CRISPR/Cas9-edited and control podocytes. Changes in both total protein levels and in phosphorylation status per site were observed. Analysis of our candidate proteins suggests that multiple signaling pathways are impaired in FD.

Abstract Recent advances in CRISPR/Cas9 technology allowing precise genome editing at a site of interest have enabled relevant human disease modeling and the development of corrective gene therapies for various genetic disorders. Paroxysmal nocturnal hemoglobinuria (PNH) is a hematological disorder linked to acquired somatic loss-of-function mutations disrupting the X-linked PIG-A gene in hematopoietic stem and progenitor cells (HSPC), characterized by the clinical triad of intravascular hemolysis, thrombosis and bone marrow failure, as well as clonal expansion of HSPC defective in glycosylphophatidylinositol(GPI)-linked proteins, due to loss of PIG-A enzymatic activity. There have been many attempts to model PNH utilizing conditional knockout strategies in mice, but these do not recapitulate the hemolytic phenotype or clonal dominance. Whether clonal expansion is intrinsic to PIG-A deficient HSPC or extrinsic, resulting from relative protection of PIG-A deficient HSPC and progeny cells from immune attack, is unclear. To explore the pathophysiologic mechanisms of PNH HSPC clonal expansion, we generated a relevant non-human primate model for PNH, utilizing autologous transplantation of HSPC edited with CRISPR/Cas9 at the PIG-A locus. The most efficient guide RNAs (gRNAs) targeting several sites within PIG-A exon 2 locus on the X chromosome, where most of mutations occur in patients with PNH, were selected based on efficiency of editing as screened in FRhK-4 cell lines and mobilized macaque CD34+ HSPC. Compared to editing of the control AAVS1 locus and other genes of interest, editing of all PIG-A locus sites was relatively inefficient. Autologous HSPC were transduced with ribonucleotide protein (RNP) complexes of Cas9 protein and selected gRNAs targeting either PIG-A or the AAVS1 "safe harbor" site, as an internal control, followed by transplantation into one female (ZI35) and one male (ZL19) macaque, respectively. Both animals engrafted promptly and clones acquiring the PNH loss-of-function phenotype, defined by loss of binding of FLAER, a fluorescent compound that binds to all GPI anchors, and loss of expression of lineage-specific GPI-linked proteins (CD24 for granulocytes and CD14 for monocytes) were stably maintained at levels of 0.2-0.4% for up to 19 months post-transplantation, with no evidence for intrinsic clonal expansion of PIG-A edited cells (Fig. 1A). Upon targeted deep sequencing, 0.4-1% insertions and deletions (INDELs) induced by CRISPR/Cas9 were detected, and the predominant INDEL type was identified as a single base deletion at the +1 or -1 positions of the target site consistently in granulocytes and other mature hematopoietic lineage cells from peripheral blood (PB), suggesting that initial mutations occurred in long-lasting HSPCs rather than short-term progenitor cells. Furthermore, as the PIG-A gene is located on X chromosome, we sought to investigate the difference in genome editing efficiency depending on the number of X alleles or activation state. Note that the expected INDEL frequency in completely PIG-A deficient sorted FLAERneg cells would be 100%, however, large deletions or rearrangements are not detected by standard deep sequencing methodologies. Interestingly, the mutation frequencies in total granulocytes and more importantly in sorted FLAER negative PNH cells were always much higher in male (ZL19) macaque cells than in female (ZI35) macaque cells (Fig. 1B and C). Consistently, gene editing PIG-A allele efficiency with CRISPR/Cas9 was also higher in human male B-lymphoblastoid cell lines (LCL) compared to female cells, 23.2% versus 16.6% (n=5), respectively. The finding that edited allele frequency was consistently lower in sorted FLAERneg female than male cells suggests that editing of the active X allele may be favored, potentially due to poor accessibility of inactive loci to editing machinery. In conclusion, we have successfully established a rhesus macaque model for PNH utilizing autologous transplantation of CRISPR/Cas9 edited HSPC. To date, we found no evidence for intrinsic expansion of PIG-A deficiency HSPC and hematopoietic progeny. This modeling approach could be utilized for further investigation of extrinsic or intrinsic factor responsible for clonal expansion. Furthermore, our findings provide a better insight into the relationship between CRISPR/Cas9 editing efficiency and active versus inactive X-linked genes. Figure 1. Figure 1. Disclosures Dunbar: National Institute of Health: Research Funding.

Abstract Objectives Neurodegeneration with brain iron accumulation (NBIA) is a clinically and genetically heterogeneous group of neurodegenerative diseases characterized by an abnormal accumulation of brain iron and progressive degeneration of the nervous system. β-propeller protein-associated neurodegeneration (BPAN) (OMIM #300,894) is a recently identified subtype of NBIA. BPAN is caused by de novo mutations in the WD repeat domain 45 (WDR45) gene. WDR45 deficiency in BPAN patients and animal models has shown defects in autophagic flux, suggesting a role for WDR45 in autophagy. How WDR45 deficiency leads to brain iron overload remains unclear. The goal of the present study is to identify the pathogenic mechanisms of WDR45 deficiency that cause iron overload and neurodegeneration. Methods To elucidate the role of WDR45 in dopaminergic neuronal cells, we generated a WDR45-knockout (KO) SH-SY5Y cell line by CRISPR/Cas9-mediated genome editing. To identify mechanisms underlying iron homeostasis and transport, we examined two cellular iron acquisition pathways in these cells using radioactive isotope 59Fe: 1) the canonical transferrin-bound iron (TBI) uptake pathway and 2) the nontransferrin-bound iron (NTBI) pathway. Results Loss of WDR45 increased total iron levels with a concomitant increase in the iron storage protein ferritin in neuronal cells. Specifically, WDR45-KO cells preferentially took up NTBI compared to wild-type cells. Concordant with these functional data, the level of divalent metal transporter-1 (DMT1) expression was upregulated in WDR45-KO cells, providing a causal link to iron overload in WDR45 deficiency. In addition, loss of WDR45 led to defects in autophagic flux and impaired ferritinophagy, a lysosomal process that promotes ferritin degradation, suggesting that iron overload is driven by impaired ferritinophagy. Interestingly, WDR45 deficiency increased iron accumulation in the mitochondria, impaired mitochondrial function, and in turn, elevated reactive oxygen species generation. Conclusions Our study provides the first evidence that WDR45 deficiency alters cellular iron acquisition pathways thereby leading to iron accumulation in neuronal cells. These findings will serve as a basis for developing therapeutic strategies for patients with NBIA. Funding Sources NIH, NBIA Disorder Association.

Abstract Background Disruptions of SETBP1 (SET binding protein 1) on 18q12.3 by heterozygous gene deletion or loss-of-function variants cause SETBP1 disorder. Clinical features are frequently associated with moderate to severe intellectual disability, autistic traits and speech and motor delays. Despite the association of SETBP1 with neurodevelopmental disorders, little is known about its role in brain development. Methods Using CRISPR/Cas9 genome editing technology, we generated a SETBP1 deletion model in human embryonic stem cells (hESCs) and examined the effects of SETBP1-deficiency in neural progenitors (NPCs) and neurons derived from these stem cells using a battery of cellular assays, genome-wide transcriptomic profiling and drug-based phenotypic rescue. Results Neural induction occurred efficiently in all SETBP1 deletion models as indicated by uniform transition into neural rosettes. However, SETBP1-deficient NPCs exhibited an extended proliferative window and a decrease in neurogenesis coupled with a deficiency in their ability to acquire ventral forebrain fate. Genome-wide transcriptome profiling and protein biochemical analysis revealed enhanced activation of Wnt/β-catenin signaling in SETBP1 deleted cells. Crucially, treatment of the SETBP1-deficient NPCs with a small molecule Wnt inhibitor XAV939 restored hyper canonical β-catenin activity and restored both cortical and MGE neuronal differentiation. Limitations The current study is based on analysis of isogenic hESC lines with genome-edited SETBP1 deletion and further studies would benefit from the use of patient-derived iPSC lines that may harbor additional genetic risk that aggravate brain pathology of SETBP1 disorder. Conclusions We identified an important role for SETBP1 in controlling forebrain progenitor expansion and neurogenic differentiation. Our study establishes a novel regulatory link between SETBP1 and Wnt/β-catenin signaling during human cortical neurogenesis and provides mechanistic insights into structural abnormalities and potential therapeutic avenues for SETBP1 disorder.

Mutations in the X-linked cyclin-dependent kinase-like 5 (CDKL5) gene cause CDKL5 deficiency disorder (CDD), which is a rare neurodevelopmental disease characterized by severe epilepsy and global developmental delay. Most children affected suffer from seizures beginning in the first months of life and severe impairment of cognitive and motor skills, with great impact on their quality of life. Most cannot walk, talk, or feed themselves, and many are confined to using a wheelchair. Although rare, CDKL5 deficiency disorder is one of the most common forms of genetic epilepsy. Currently, there is no cure or effective treatment for CDD, hence the great urge to develop novel and effective therapeutic strategies. Here, we present a methodology for the correction of a pathogenic variant in CDKL5 (c.1090G>T (p.Glu364*)), using CRISPR/Cas9 genome editing technology in patient-derived cell models, in order to expedite the discovery of new therapies for CDD. CRISPR/Cas9 is a precise and versatile method of genetic manipulation, and it only requires three components to target and correct genetic mutations: guide RNAs, Cas9 endonuclease, and homology-directed repair (HDR) templates. We first tested plasmid-based delivery of CRISPR/Cas9 for correction in primary fibroblasts. This system proved to be up to 66% efficient but it was associated with extremely variable and unpredictable editing efficiency (33±31%) in three separate experiments. Hence, we decided to test additional guides and to replace the plasmid-based system with a protein-based ribonucleoprotein (RNP) delivery system for more rapid action and greater stability. We tested the system in induced pluripotent stem cells (iPSCs) obtained by reprogramming the patient’s fibroblasts. We reported the generation of genetically corrected iPSCs, where the mutated CDKL5 c.1090G>T (p.Glu364*) was corrected to the wild-type, using RNP-mediated delivery of CRISPR/Cas9. Based on PCR cloning results of gene-corrected clones, we can state that our system is able to selectively target the p.Glu364* variant while preserving the wild-type CDKL5 allele in vitro. We then differentiated in parallel mutant and isogenic sets of cells into neural cells to assess the functional consequences of the edit in the affected cell type. We demonstrated that CRISPR/Cas9 gene editing restores the expression of CDKL5 protein in iPSC-derived neurons by Western blot. We also showed by RT-qPCR that mutant neurons carrying the c.1090G>T (p.Glu364*) in CDKL5 pre-sent reduced expression of CDKL5 mRNA compared to isogenic control. Our findings demonstrate that we can achieve targeted and allele-specific correction of CDKL5 (c.1090G>T (p.Glu364*)) variant using CRISPR/Cas9-RNP system in a patient-specific cell model. Moreover, we proved that correction of the mutation at the DNA level rescues CDKL5 protein expression and increases CDKL5 mRNA expression in isogenic neurons. The results of this study might be decisive in proving CRISPR/Cas9 potential to carry out genome editing in human cells, and ultimately for developing advanced therapies for CDD.