Liren Wang

Heritable gene targeting in the mouse and rat using a CRISPR-Cas system.

To the Editor: CRISPR-Cas systems have been developed as an efficient gene editing technology in cells and model organisms. Here we use a CRISPR-Cas system to induce genomic DNA fragment deletion in mice by coinjecting two single-guide RNAs (sgRNAs) targeting the Uhrf2 locus with Cas9 mRNA. Furthermore, we report the generation of a Mc3R and Mc4R double-gene knockout rat by means of a single microinjection. High germline-transmission efficiency was observed in both mice and rats. The clustered, regularly interspaced, short palindromic repeats (CRISPR)associated protein (Cas) system has evolved in bacteria and archaea as an RNA-based adaptive immune system against viral and plasmid invasion1. Based on gene conservation and locus organization, three major types of CRISPR systems have been identified2,3. In the type II systems, the complex of a CRISPR RNA (crRNA) annealed to a trans-activating crRNA (tracrRNA) is sufficient to guide the Cas9 endonuclease to a specific genomic sequence to generate double-strand breaks in target DNA4. Previous studies established a strategy for multiplex genome engineering with the Cas9 RNA-guided endonuclease in mammalian cells5,6. Recently, efficient genome editing by the CRISPR-Cas system has been shown in multiple organisms, including zebrafish, mice and bacteria7–9. Several groups have demonstrated that compared with zinc finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs), CRISPR-Cas–mediated gene targeting has similar or greater efficiency in cells and zebrafish5–7,10. Although it has been demonstrated that multiple genes can be disrupted in individual mouse embryos using CRISPR-Cas–mediated systems9, germline transmission of Cas9-mediated mutations in animals has not yet been reported. In addition, whether long, specific, genomic DNA target fragments

CRISPR/Cas-mediated genome editing in the rat via direct injection of one-cell embryos

Conventional embryonic stem cell (ESC)–based gene targeting, zinc-finger nuclease (ZFN) and transcription activator–like effector nuclease (TALEN) technologies are powerful strategies for the generation of genetically modified animals. Recently, the CRISPR/Cas system has emerged as an efficient and convenient alternative to these approaches. We have used the CRISPR/Cas system to generate rat strains that carry mutations in multiple genes through direct injection of RNAs into one-cell embryos, demonstrating the high efficiency of Cas9-mediated gene editing in rats for simultaneous generation of compound gene mutant models. Here we describe a stepwise procedure for the generation of knockout and knock-in rats. This protocol provides guidelines for the selection of genomic targets, synthesis of guide RNAs, design and construction of homologous recombination (HR) template vectors, embryo microinjection, and detection of mutations and insertions in founders or their progeny. The procedure from target design to identification of founders can take as little as 6 weeks, of which <10 d is actual hands-on working time.

Large genomic fragment deletion and functional gene cassette knock-in via Cas9 protein mediated genome editing in one-cell rodent embryos

The CRISPR-Cas RNA-guided system has versatile uses in many organisms and allows modification of multiple target sites simultaneously. Generating novel genetically modified mouse and rat models is one valuable application of this system. Through the injection of Cas9 protein instead of mRNA into embryos, we observed fewer off-target effects of Cas9 and increased point mutation knock-in efficiency. Large genomic DNA fragment (up to 95 kb) deletion mice were generated for in vivo study of lncRNAs and gene clusters. Site-specific insertion of a 2.7 kb CreERT2 cassette into the mouse Nfatc1 locus allowed labeling and tracing of hair follicle stem cells. In addition, we combined the Cre-Loxp system with a gene-trap strategy to insert a GFP reporter in the reverse orientation into the rat Lgr5 locus, which was later inverted by Cre-mediated recombination, yielding a conditional knockout/reporter strategy suitable for mosaic mutation analysis.

CRISPR/Cas9‐mediated somatic correction of a novel coagulator factor IX gene mutation ameliorates hemophilia in mouse

The X‐linked genetic bleeding disorder caused by deficiency of coagulator factor IX, hemophilia B, is a disease ideally suited for gene therapy with genome editing technology. Here, we identify a family with hemophilia B carrying a novel mutation, Y371D, in the human F9 gene. The CRISPR/Cas9 system was used to generate distinct genetically modified mouse models and confirmed that the novel Y371D mutation resulted in a more severe hemophilia B phenotype than the previously identified Y371S mutation. To develop therapeutic strategies targeting this mutation, we subsequently compared naked DNA constructs versus adenoviral vectors to deliver Cas9 components targeting the F9 Y371D mutation in adult mice. After treatment, hemophilia B mice receiving naked DNA constructs exhibited correction of over 0.56% of F9 alleles in hepatocytes, which was sufficient to restore hemostasis. In contrast, the adenoviral delivery system resulted in a higher corrective efficiency but no therapeutic effects due to severe hepatic toxicity. Our studies suggest that CRISPR/Cas‐mediated in situ genome editing could be a feasible therapeutic strategy for human hereditary diseases, although an efficient and clinically relevant delivery system is required for further clinical studies.

PAK1IP1, a ribosomal stress-induced nucleolar protein, regulates cell proliferation via the p53–MDM2 loop

Cell growth and proliferation are tightly controlled via the regulation of the p53–MDM2 feedback loop in response to various cellular stresses. In this study, we identified a nucleolar protein called PAK1IP1 as another regulator of this loop. PAK1IP1 was induced when cells were treated with chemicals that disturb ribosome biogenesis. Overexpression of PAK1IP1 inhibited cell proliferation by inducing p53-dependent G1 cell-cycle arrest. PAK1IP1 bound to MDM2 and inhibited its ability to ubiquitinate and to degrade p53, consequently leading to the accumulation of p53 levels. Interestingly, knockdown of PAK1IP1 in cells also inhibited cell proliferation and induced p53-dependent G1 arrest. Deficiency of PAK1IP1 increased free ribosomal protein L5 and L11 which were required for PAK1IP1 depletion-induced p53 activation. Taken together, our results reveal that PAK1IP1 is a new nucleolar protein that is crucial for rRNA processing and plays a regulatory role in cell proliferation via the p53–MDM2 loop.

Characterization of novel cytochrome P450 2E1 knockout rat model generated by CRISPR/Cas9.

A bacterial CRISPR-associated protein-9 nuclease (CRISPR/Cas9) from Streptococcus pyogenes has generated considerable excitement as a new tool to edit the targeted genome. Cytochrome P450 (CYP) 2E1 not only plays an important role in the xenobiotic metabolism and chemical toxicity, but also is involved in many kinds of diseases, such as alcoholic liver diseases and diabetes. Despite its importance, few animal models are used to predict CYP2E1 properties in physiology, pathology, as well as carcinogen activation. To establish a novel model for investigating the functions of CYP2E1 in vivo, this study has successfully generated the Cyp2e1 knockout (KO) rat model without detectable off-target effects using CRISPR/Cas9 system. The Cyp2e1 KO rats were viable and fertile and did not display any obvious physiological abnormities. The absent expression of CYP2E1 in KO rats also resulted in inactive behaviors in the metabolism of CYP2E1 substrates. The Cyp2e1 KO rats as a novel and available rodent animal model provide a powerful tool for the study of CYP2E1 in the chemical metabolism, toxicity, carcinogenicity, and its core factor in drug-drug interactions.

Increasing targeting scope of adenosine base editors in mouse and rat embryos through fusion of TadA deaminase with Cas9 variants

The clustered regularly interspaced short palindromic repeat (CRISPR) system has been widely adapted to genome editing to either introduce or correct genetic mutations (Wang et al., 2016). However, due to competition with the dominant non-homologous end-joining (NHEJ) pathway, precise genome modifications through Cas9-stimulated homologous recombination (HR) is inefficient. Through fusion of cytidine deaminases, APOBEC1 (apolipoprotein B editing complex 1) or AID (activation-induced deaminase), with Cas9 variants, several groups have developed the cytidine base editor (BE) systems (Komor et al., 2016; Li et al., 2018; Nishida et al., 2016). The BE system achieves programmable conversion of C•G base pairs to T•A without double-stranded DNA cleavage (Zhou et al., 2017). More recently, adenine base editors (ABEs), which efficiently convert A•T base pairs to G•C in genomic DNA, have been developed via fusion of an engineered tRNA adenosine deaminase (ecTadA from Escherichia coli) with nCas9 (Gaudelli et al., 2017). The ABE system has quickly been adapted to generate disease models and correction of genetic disease in mice (Ryu et al., 2017; Liu et al., 2018). However, whether the editing efficiency and the targeting scope of ABE could be improved is largely unexplored. In this study, we describe the efficient generation of base-edited mice and rats modeling human diseases through ABEs with highest efficiency up to 100%. We also demonstrate an increase of ABE activity through injection of chemically modified tracrRNA and crRNA in mouse zygotes, and the expansion of editing scope by fusion of an ecTadA mutant to SaCas9n-KKH and Cas9n-VQR variants in both cells and embryos. Our study suggests that the ABE system is a powerful and convenient tool to introduce precise base conversions in rodents. To test the ABE efficiency in embryos, we injected ABE mRNA (Fig. 1A) together with sgRNA targeting the TATA box of the Hbb-bs gene, into C57BL6 strain mouse zygotes (Fig. S1A and Table S1). Overlapping A/G peaks in the target sites were identified in 14/27 of F0 mice as determined by the chromatograms of Sanger sequencing (Figs. 2F and S1B). Further analysis by deep sequencing revealed allelic frequencies from 6%–71% among the founders (Fig. S1C). In individual allele, the editing window was extended from position A2–A9 in mouse embryos, which is broader than the window spanning position A4–A7 observed in mammalian cell lines (Gaudelli et al., 2017) (Fig. S1B and S1C). These data demonstrate that ABE is efficient to generate point mutant mice and its mutation window expands in embryos. Next, we tested the capability of ABE to precisely mutate A:T pairs for disrupting the stop codon of the gene encoding the fumarylacetoacetate hydrolase (Fah) (Fig. 1B), whose mutations cause hereditary tyrosinemia type I (HTI) in humans. We observed high A>G conversion efficiency (39/47) among F0 mice with allelic frequencies varying from 7%–99% as determined by deep sequencing (Figs. 1B, 1C, 2F and S2A). Increasing the sgRNA concentration from 50 ng/μL to 100 ng/μL results in 100% (13/13) point mutation rate in F0 mice (Figs. 2F and S2B). Since disruption of the stop codon usually affects mRNA stability and protein expression (Frischmeyer et al., 2002), the Fah mRNA and protein levels were dramatically impaired (Fig. S2C and S2D). Through immunohistochemistry analysis of the liver tissue from founder F0–F32, Fah protein expression was almost undetectable (Fig. 1D) suggesting this founder was a homozygote (Fig. 1B and 1C). To investigate germline transmission efficiency, founder mice were crossed with wild type or with other founders. We observed high germline transmission efficiency (Fig. S3A and S3B). In homozygous F1 mice, the expression of Fah mRNA and protein was lost (Fig. S3C–E). After withdrawal of the 2-(2-nitro-4-trifluoromethylbenzoyl)-1,3-cyclohexanedione (NTBC) treatment, the phenotypes of Fah mutant homozygotes were similar to previous HTI model, including loss of body weight and perturbation of serum biomarkers (Shao et al., 2018) (Fig. S3F and S3G). As mutations that generate premature stop codons are common drivers in various genetic diseases (Keeling et al., 2014), ABE has a promising potential for readthrough of premature stop codons in certain genetic diseases as demonstrated in the mouse DMD model (Ryu et al., 2017). Previous study demonstrated that 2’-O-methyl-3’-phosphorothioate (MS) modification on each ends of RNA can increase its stability, thus enhancing the Cas9 genome

Cas9-nickase–mediated genome editing corrects hereditary tyrosinemia in rats

Hereditary tyrosinemia type I (HTI) is a metabolic genetic disorder caused by mutation of fumarylacetoacetate hydrolase (FAH). Because of the accumulation of toxic metabolites, HTI causes severe liver cirrhosis, liver failure, and even hepatocellular carcinoma. HTI is an ideal model for gene therapy, and several strategies have been shown to ameliorate HTI symptoms in animal models. Although CRISPR/Cas9-mediated genome editing is able to correct the Fah mutation in mouse models, WT Cas9 induces numerous undesired mutations that have raised safety concerns for clinical applications. To develop a new method for gene correction with high fidelity, we generated a Fah mutant rat model to investigate whether Cas9 nickase (Cas9n)-mediated genome editing can efficiently correct the Fah. First, we confirmed that Cas9n rarely induces indels in both on-target and off-target sites in cell lines. Using WT Cas9 as a positive control, we delivered Cas9n and the repair donor template/single guide (sg)RNA through adenoviral vectors into HTI rats. Analyses of the initial genome editing efficiency indicated that only WT Cas9 but not Cas9n causes indels at the on-target site in the liver tissue. After receiving either Cas9n or WT Cas9-mediated gene correction therapy, HTI rats gained weight steadily and survived. Fah-expressing hepatocytes occupied over 95% of the liver tissue 9 months after the treatment. Moreover, CRISPR/Cas9-mediated gene therapy prevented the progression of liver cirrhosis, a phenotype that could not be recapitulated in the HTI mouse model. These results strongly suggest that Cas9n-mediated genome editing is a valuable and safe gene therapy strategy for this genetic disease.