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Cut and Paste: Efficient Homology-Directed Repair of a Dominant Negative KRT14 Mutation via CRISPR/Cas9 Nickases.
With the ability to induce rapid and efficient repair of disease-causing mutations, CRISPR/Cas9 technology is ideally suited for gene therapy approaches for recessively and dominantly inherited monogenic disorders. In this study, we have corrected a causal hotspot mutation in exon 6 of the keratin 14 gene (KRT14) that results in generalized severe epidermolysis bullosa simplex (EBS-gen sev), using a double-nicking strategy targeting intron 7, followed by homology-directed repair (HDR). Co-delivery into EBS keratinocytes of a Cas9 D10A nickase (Cas9n), a predicted single guide RNA pair specific for intron 7, and a minicircle donor vector harboring the homology donor template resulted in a recombination efficiency of >30% and correction of the mutant KRT14 allele. Phenotypic correction of EBS-gen sev keratinocytes was demonstrated by immunofluorescence analysis, revealing the absence of disease-associated K14 aggregates within the cytoplasm. We achieved a promising safety profile for the CRISPR/Cas9 double-nicking approach, with no detectable off-target activity for a set of predicted off-target genes as confirmed by next generation sequencing. In conclusion, we demonstrate a highly efficient and specific gene-editing approach for KRT14, offering a causal treatment option for EBS.
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Engineering a precise adenine base editor with minimal bystander editing.
Adenine base editors (ABEs) catalyze A-to-G transitions showing broad applications, but their bystander mutations and off-target editing effects raise safety concerns. Through structure-guided engineering, we found ABE8e with an N108Q mutation reduced both adenine and cytosine bystander editing, and introduction of an additional L145T mutation (ABE9), further refined the editing window to 1-2 nucleotides with eliminated cytosine editing. Importantly, ABE9 induced very minimal RNA and undetectable Cas9-independent DNA off-target effects, which mainly installed desired single A-to-G conversion in mouse and rat embryos to efficiently generate disease models. Moreover, ABE9 accurately edited the A5 position of the protospacer sequence in pathogenic homopolymeric adenosine sites (up to 342.5-fold precision over ABE8e) and was further confirmed through a library of guide RNA-target sequence pairs. Owing to the minimized editing window, ABE9 could further broaden the targeting scope for precise correction of pathogenic single-nucleotide variants when fused to Cas9 variants with expanded protospacer adjacent motif compatibility. bpNLS, bipartite nuclear localization signals.
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CRISPR C-to-G base editors for inducing targeted DNA transversions in human cells.
CRISPR-guided DNA cytosine and adenine base editors are widely used for many applications1-4 but primarily create DNA base transitions (that is, pyrimidine-to-pyrimidine or purine-to-purine). Here we describe the engineering of two base editor architectures that can efficiently induce targeted C-to-G base transversions, with reduced levels of unwanted C-to-W (W = A or T) and indel mutations. One of these C-to-G base editors (CGBE1), consists of an RNA-guided Cas9 nickase, an Escherichia coli-derived uracil DNA N-glycosylase (eUNG) and a rat APOBEC1 cytidine deaminase variant (R33A) previously shown to have reduced off-target RNA and DNA editing activities5,6. We show that CGBE1 can efficiently induce C-to-G edits, particularly in AT-rich sequence contexts in human cells. We also removed the eUNG domain to yield miniCGBE1, which reduced indel frequencies but only modestly decreased editing efficiency. CGBE1 and miniCGBE1 enable C-to-G edits and will serve as a basis for optimizing C-to-G base editors for research and therapeutic applications.
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Genome editing approaches to β-hemoglobinopathies.
β-hemoglobinopathies are the most common monogenic disorders worldwide and are caused by mutations in the β-globin locus altering the production of adult hemoglobin (HbA). Transplantation of autologous hematopoietic stem cells (HSCs) corrected by lentiviral vector-mediated addition of a functional β-like globin raised new hopes to treat sickle cell disease and β-thalassemia patients; however, the low expression of the therapeutic gene per vector copy is often not sufficient to fully correct the patients with a severe clinical phenotype. Recent advances in the genome editing field brought new possibilities to cure β-hemoglobinopathies by allowing the direct modification of specific endogenous loci. Double-strand breaks (DSBs)-inducing nucleases (i.e., ZFNs, TALENs and CRISPR-Cas9) or DSB-free tools (i.e., base and prime editing) have been used to directly correct the disease-causing mutations, restoring HbA expression, or to reactivate the expression of the fetal hemoglobin (HbF), which is known to alleviate clinical symptoms of β-hemoglobinopathy patients. Here, we describe the different genome editing tools, their application to develop therapeutic approaches to β-hemoglobinopathies and ongoing clinical trials using genome editing strategies.
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Generation and propagation of high fecundity gene edited fine wool sheep by CRISPR/Cas9.
CRISPR/Cas9 technology has been widely utilized to enhance productive performance, increase disease resistance and generate medical models in livestock. The FecB allele in sheep is a mutation in the BMPRIB gene, recognized as the first major gene responsible for the high fecundity trait in sheep, leading to an increased ovulation rate in ewe. In this study, we employed CRISPR/Cas9-mediated homologous-directed repair (HDR) to introduce a defined point mutation (c.746 A > G) using single-stranded oligonucleotides (ssODN) and the ligase IV inhibitor (SCR7) into the BMPRIB gene of fine wool sheep. A total of nine gene-edited sheep were produced, six of which carried the targeted point mutation, with a precise base substitution efficiency (A > G) of 31.6%. Based on the six targeted founders (F0), we expanded the BMPRIB-targeted population, which included F1 heterozygous (B+) and F2 homozygous(BB) or heterozygous offspring. The average litter size of F1 ewes carrying the B + allele reached 170%, comparable to that of heterozygous native Australian Booroola sheep. Gene-edited ewes with B + and BB genotype produced 0.62 and 0.42 more lambs, respectively, compared to wide-type ewes (p < 0.01). Our results also indicated that the parity signification, our data demonstrate that highly efficient introduction of the intended base mutation into the sheep genome can be achieved by combining the CRISPR/Cas9 system with ssODN and SCR7. The offspring of BMPR/B edited sheep with the defined mutation exhibited high fecundity performance. Compared to conventional sheep breeding strategies, genetic improvement through gene editing offered significant advantages without compromising the fine wool traits of Merino sheep, which are often affected by routine cross-breeding methods.
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Efficient Generation and Correction of Mutations in Human iPS Cells Utilizing mRNAs of CRISPR Base Editors and Prime Editors.
In contrast to CRISPR/Cas9 nucleases, CRISPR base editors (BE) and prime editors (PE) enable predefined nucleotide exchanges in genomic sequences without generating DNA double strand breaks. Here, we employed BE and PE mRNAs in conjunction with chemically synthesized sgRNAs and pegRNAs for efficient editing of human induced pluripotent stem cells (iPSC). Whereas we were unable to correct a disease-causing mutation in patient derived iPSCs using a CRISPR/Cas9 nuclease approach, we corrected the mutation back to wild type with high efficiency utilizing an adenine BE. We also used adenine and cytosine BEs to introduce nine different cancer associated TP53 mutations into human iPSCs with up to 90% efficiency, generating a panel of cell lines to investigate the biology of these mutations in an isogenic background. Finally, we pioneered the use of prime editing in human iPSCs, opening this important cell type for the precise modification of nucleotides not addressable by BEs and to multiple nucleotide exchanges. These approaches eliminate the necessity of deriving disease specific iPSCs from human donors and allows the comparison of different disease-causing mutations in isogenic genetic backgrounds.
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A simple, flexible and high-throughput cloning system for plant genome editing via CRISPR-Cas system.
CRISPR-Cas9 system is now widely used to edit a target genome in animals and plants. Cas9 protein derived from Streptococcus pyogenes (SpCas9) cleaves double-stranded DNA targeted by a chimeric single-guide RNA (sgRNA). For plant genome editing, Agrobacterium-mediated T-DNA transformation has been broadly used to express Cas9 proteins and sgRNAs under the control of CaMV 35S and U6/U3 promoter, respectively. We here developed a simple and high-throughput binary vector system to clone a 19-20 bp of sgRNA, which binds to the reverse complement of a target locus, in a large T-DNA binary vector containing an SpCas9 expressing cassette. Two-step cloning procedures: (1) annealing two target-specific oligonucleotides with overhangs specific to the AarI restriction enzyme site of the binary vector; and (2) ligating the annealed oligonucleotides into the two AarI sites of the vector, facilitate the high-throughput production of the positive clones. In addition, Cas9-coding sequence and U6/U3 promoter can be easily exchanged via the Gateway(TM) system and unique EcoRI/XhoI sites on the vector, respectively. We examined the mutation ratio and patterns when we transformed these constructs into Arabidopsis thaliana and a wild tobacco, Nicotiana attenuata. Our vector system will be useful to generate targeted large-scale knock-out lines of model as well as non-model plant.
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CRISPR technology: A versatile tool to model, screen, and reverse drug resistance in cancer.
Drug resistance is a serious challenge in cancer treatment that can render chemotherapy a failure. Understanding the mechanisms behind drug resistance and developing novel therapeutic approaches are cardinal steps in overcoming this issue. Clustered regularly interspaced short palindrome repeats (CRISPR) gene-editing technology has proven to be a useful tool to study cancer drug resistance mechanisms and target the responsible genes. In this review, we evaluated original research studies that used the CRISPR tool in three areas related to drug resistance, namely screening resistance-related genes, generating modified models of resistant cells and animals, and removing resistance by genetic manipulation. We reported the targeted genes, study models, and drug groups in these studies. In addition to discussing different applications of CRISPR technology in cancer drug resistance, we analyzed drug resistance mechanisms and provided examples of CRISPR's role in studying them. Although CRISPR is a powerful tool for examining drug resistance and sensitizing resistant cells to chemotherapy, more studies are required to overcome its disadvantages, such as off-target effects, immunotoxicity, and inefficient delivery of CRISPR/cas9 into the cells.
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The case of the missing Ks: Base editor screen to assess cellular fitness at single lysines.
In this issue of Molecular Cell, Bao et al.1 set out to elucidate "functional lysines" in the genome using adenine base editors. The study reveals several cases of alteration of functions that previous canonical CRISPR-Cas9 screens were unable to detect.
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Targeted mutagenesis in Arabidopsis thaliana using CRISPR-Cas12b/C2c1.
Cas12b/C2c1 is a newly identified class 2 CRISPR endonuclease that was recently engineered for targeted genome editing in mammals and rice. To explore the potential applications of the CRISPR-Cas12b system in the dicot Arabidopsis thaliana, we selected BvCas12b and BhCas12b v4 for analysis. We successfully used both endonucleases to induce mutations, perform multiplex genome editing, and create large deletions at multiple loci. No significant mutations were detected at potential off-target sites. Analysis of the insertion/deletion frequencies and patterns of mutants generated via targeted gene mutagenesis highlighted the potential utility of CRISPR-Cas12b systems for genome editing in Arabidopsis.
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