Removal of the GAA repeat in the heart of a Friedreich's ataxia mouse model using CjCas9.

Most Friedreich ataxia (FRDA) cases are caused by the elongation of the GAA repeat (GAAr) sequence in the first intron of the FXN gene, leading to a decrease of the frataxin protein expression. Deletion of this GAAr with CRISPR/Cas9 technology leads to an increase in frataxin expression in vitro. We are therefore aiming to develop FRDA treatment based on the deletion of GAAr with CRISPR/Cas9 technology using a single AAV expressing a small Cas9 (CjCas9) and two single guide RNAs (sgRNAs) targeting the FXN gene. This AAV was intraperitoneally administrated to YG8sR (250-300 GAAr) and to YG8-800 (800 GAAr) mice. DNA and RNA were extracted from different organs a month later. PCR amplification of part of intron 1 of the FXN gene detected some GAAr deletion in some cells in heart and liver of both mouse models, but the editing rate was not sufficient to cause an increase in frataxin mRNA in the heart. However, the correlation observed between the editing rate and the distribution of AAV suggests a possible therapy based on the removal of the GAAr with a better delivery tool of the CRISPR/Cas9 system.

Serum extracellular vesicles for delivery of CRISPR-CAS9 ribonucleoproteins to modify the dystrophin gene.

Extracellular vesicles (EVs) mediate intercellular biomolecule exchanges in the body, making them promising delivery vehicles for therapeutic cargo. Genetic engineering by the CRISPR system is an interesting therapeutic avenue for genetic diseases such as Duchenne muscular dystrophy (DMD). We developed a simple method for loading EVs with CRISPR ribonucleoproteins (RNPs) consisting of SpCas9 proteins and guide RNAs (gRNAs). EVs were first purified from human or mouse serum using ultrafiltration and size-exclusion chromatography. Using protein transfectant to load RNPs into serum EVs, we showed that EVs are good carriers of RNPs in vitro and restored the expression of the tdTomato fluorescent protein in muscle fibers of Ai9 mice. EVs carrying RNPs targeting introns 22 and 24 of the DMD gene were also injected into muscles of mdx mice having a non-sense mutation in exon 23. Up to 19% of the cDNA extracted from treated mdx mice had the intended deletion of exons 23 and 24, allowing dystrophin expression in muscle fibers. RNPs alone, without EVs, were inefficient in generating detectable deletions in mouse muscles. This method opens new opportunities for rapid and safe delivery of CRISPR components to treat DMD.

CRISPR-SCReT (CRISPR-Stop Codon Read Through) method to control Cas9 expression for gene editing.

CRISPR/Cas9 has paved the way for the development of therapies that correct genetic mutations. However, constitutive expression of the Cas9 gene can increase off-target mutations and induce an immune response against the Cas9 protein. To limit the time during which the Cas9 nuclease is expressed, we proposed a simple drug inducible system. The approach consists of introducing a premature termination codon (PTC) in the Cas9 gene and subsequently treating with an aminoglycoside drug, which allows readthrough of the complete protein. To validate that system, HEK293T cells were co-transfected with a PX458 plasmid, which was mutated to introduce a PTC in the SpCas9 gene and two sgRNAs targeting the DMD gene (exons 50 and 54). Cells were treated with different doses of geneticin (G418) for 48 h. Western blot confirmed that the Cas9 protein expression, which was shut down by the PTC mutation, can be induced by the drug. The hybrid exon 50-54 formed by the deletion of part of the DMD gene was detected by PCR only in the cells treated with G418. The approach was also used successfully with CjCas9 to edit the FXN gene. Our results show that it is possible to control SpCas9 and CjCas9 expression by CRISPR-SCReT (CRISPR-Stop Codon Read Through) method.

Current Clinical Applications of In Vivo Gene Therapy with AAVs.

Hereditary diseases are caused by mutations in genes, and more than 7,000 rare diseases affect over 30 million Americans. For more than 30 years, hundreds of researchers have maintained that genetic modifications would provide effective treatments for many inherited human diseases, offering durable and possibly curative clinical benefit with a single treatment. This review is limited to gene therapy using adeno-associated virus (AAV) because the gene delivered by this vector does not integrate into the patient genome and has a low immunogenicity. There are now five treatments approved for commercialization and currently available, i.e., Luxturna, Zolgensma, the two chimeric antigen receptor T cell (CAR-T) therapies (Yescarta and Kymriah), and Strimvelis (the gammaretrovirus approved for adenosine deaminase-severe combined immunodeficiency [ADA-SCID] in Europe). Dozens of other treatments are under clinical trials. The review article presents a broad overview of the field of therapy by in vivo gene transfer. We review gene therapy for neuromuscular disorders (spinal muscular atrophy [SMA]; Duchenne muscular dystrophy [DMD]; X-linked myotubular myopathy [XLMTM]; and diseases of the central nervous system, including Alzheimer's disease, Parkinson's disease, Canavan disease, aromatic l-amino acid decarboxylase [AADC] deficiency, and giant axonal neuropathy), ocular disorders (Leber congenital amaurosis, age-related macular degeneration [AMD], choroideremia, achromatopsia, retinitis pigmentosa, and X-linked retinoschisis), the bleeding disorder hemophilia, and lysosomal storage disorders.

Prime Editing Permits the Introduction of Specific Mutations in the Gene Responsible for Duchenne Muscular Dystrophy.

The Prime editing technique derived from the CRISPR/Cas9 discovery permits the modification of selected nucleotides in a specific gene. We used it to insert specific point mutations in exons 9, 20, 35, 43, 55 and 61 of the Duchenne Muscular Dystrophy (DMD) gene coding for the dystrophin protein, which is absent in DMD patients. Up to 11% and 21% desired mutations of the DMD gene in HEK293T cells were obtained with the PRIME Editor 2 (PE2) and PE3, respectively. Three repeated treatments increased the percentage of specific mutations with PE2 to 16%. An additional mutation in the protospacer adjacent motif (PAM) sequence improved the PE3 result to 38% after a single treatment. We also carried out the correction of c.428 G>A point mutation in exon 6 of the DMD gene in a patient myoblast. Myoblast electroporation showed up to 8% and 28% modifications, respectively, for one and three repeated treatments using the PE3 system. The myoblast correction led to dystrophin expression in myotubes detected by Western blot. Thus, prime editing can be used for the correction of point mutations in the DMD gene.

CRISPR-Induced Deletion with SaCas9 Restores Dystrophin Expression in Dystrophic Models In Vitro and In Vivo.

Duchenne muscular dystrophy (DMD), a severe hereditary disease affecting 1 in 3,500 boys, mainly results from the deletion of exon(s), leading to a reading frameshift of the DMD gene that abrogates dystrophin protein synthesis. Pairs of sgRNAs for the Cas9 of Staphylococcus aureus were meticulously chosen to restore a normal reading frame and also produce a dystrophin protein with normally phased spectrin-like repeats (SLRs), which is not usually obtained by skipping or by deletion of complete exons. This can, however, be obtained in rare instances where the exon and intron borders of the beginning and the end of the complete deletion (patient deletion plus CRISPR-induced deletion) are at similar positions in the SLR. We used pairs of sgRNAs targeting exons 47 and 58, and a normal reading frame was restored in myoblasts derived from muscle biopsies of 4 DMD patients with different exon deletions. Restoration of the DMD reading frame and restoration of dystrophin expression were also obtained in vivo in the heart of the del52hDMD/mdx. Our results provide a proof of principle that SaCas9 could be used to edit the human DMD gene and could be considered for further development of a therapy for DMD.

Nonfunctional mutant Wrn protein leads to neurological deficits, neuronal stress, microglial alteration, and immune imbalance in a mouse model of Werner syndrome.

Werner syndrome (WS) is a premature aging disorder caused by mutations in a RecQ-family DNA helicase, WRN. Mice lacking part of the helicase domain of the WRN orthologue exhibit many phenotypic features of WS, including metabolic abnormalities and a shorter lifespan. Yet, little is known about the impact of WRN mutations on the central nervous system in both humans and mouse models of WS. In the current study, we have performed a longitudinal behavioral assessment on mice bearing a Wrn helicase deletion. Behavioral tests demonstrated a loss of motor activity and coordination, reduction in perception, increase in repetitive behavior, and deficits in both spatial and social novelty memories in Wrn mutant mice compared to age-matched wild type mice. These neurological deficits were associated with biochemical and histological changes in the brain of aged Wrn mutant mice. Microglia, resident immune cells that regulate neuronal plasticity and function in the brain, were hyper-ramified in multiple regions involved with the behavioral deficits of Wrn mutant mice. Furthermore, western analyses indicated that Wrn mutant mice exhibited an increase of oxidative stress markers in the prefrontal cortex. Supporting these findings, electron microscopy studies revealed increased cellular aging and oxidative stress features, among microglia and neurons respectively, in the prefrontal cortex of aged Wrn mutant mice. In addition, multiplex immunoassay of serum identified significant changes in the expression levels of several pro- and anti-inflammatory cytokines. Taken together, these findings indicate that microglial dysfunction and neuronal oxidative stress, associated with peripheral immune system alterations, might be important driving forces leading to abnormal neurological symptoms in WS thus suggesting potential therapeutic targets for interventions.

Generation of human endometrial knockout cell lines with the CRISPR/Cas9 system confirms the prostaglandin F2α synthase activity of aldo-ketoreductase 1B1.

Prostaglandins (PGs) are important regulators of female reproductive function. The primary PGs produced in the endometrium are PGE2 and PGF2α. Relatively little is known about the biosynthetic pathways leading to the formation of PGF2α. We have described the role of aldo-ketoreductase (AKR)1B1 in increased PGF2α production by human endometrial cells following stimulation with interleukin-1ß (IL-1ß). However, alternate PGF synthases are expressed concurrently in endometrial cells. A definite proof of the role of AKR1B1 would require gene knockout; unfortunately, this gene has no direct equivalent in the mouse. Recently, an efficient genome-editing technology using RNA-guided DNase Cas9 and the clustered regularly interspaced short palindromic repeats (CRISPR) system has been developed. We have adapted this approach to knockout AKR1B1 gene expression in human endometrial cell lines. One clone (16-2) of stromal origin generated by the CRISPR/Cas9 system exhibited a complete loss of AKR1B1 protein and mRNA expression, whereas other clones presented with partial edition. The present report focuses on the characterization of clone 16-2 exhibiting deletion of 68 and 2 nucleotides, respectively, on each of the alleles. Cells from this clone lost their ability to produce PGF2α but maintained their original stromal cell (human endometrial stromal cells-2) phenotype including the capacity to decidualize in the presence of progesterone (medroxyprogesterone acetate) and 8-bromo-cAMP. Knockout cells also maintained their ability to increase PGE2 production in response to IL-1ß. In summary, we demonstrate that the new genome editing CRISPR/Cas9 system can be used in human cells to generate stable knockout cell line models. Our results suggest that genome editing of human cell lines can be used to complement mouse KO models to validate the function of genes in differentiated tissues and cells. Our results also confirm that AKR1B1 is involved in the synthesis of PGF2α.

Reading frame correction by targeted genome editing restores dystrophin expression in cells from Duchenne muscular dystrophy patients.

Genome editing with engineered nucleases has recently emerged as an approach to correct genetic mutations by enhancing homologous recombination with a DNA repair template. However, many genetic diseases, such as Duchenne muscular dystrophy (DMD), can be treated simply by correcting a disrupted reading frame. We show that genome editing with transcription activator-like effector nucleases (TALENs), without a repair template, can efficiently correct the reading frame and restore the expression of a functional dystrophin protein that is mutated in DMD. TALENs were engineered to mediate highly efficient gene editing at exon 51 of the dystrophin gene. This led to restoration of dystrophin protein expression in cells from Duchenne patients, including skeletal myoblasts and dermal fibroblasts that were reprogrammed to the myogenic lineage by MyoD. Finally, 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. This strategy integrates the rapid and robust assembly of active TALENs with an efficient gene-editing method for the correction of genetic diseases caused by mutations in non-essential coding regions that cause frameshifts or premature stop codons.