Generation of rat forebrain tissues in mice.

Interspecies blastocyst complementation (IBC) provides a unique platform to study development and holds the potential to overcome worldwide organ shortages. Despite recent successes, brain tissue has not been achieved through IBC. Here, we developed an optimized IBC strategy based on C-CRISPR, which facilitated rapid screening of candidate genes and identified that Hesx1 deficiency supported the generation of rat forebrain tissue in mice via IBC. Xenogeneic rat forebrain tissues in adult mice were structurally and functionally intact. Cross-species comparative analyses revealed that rat forebrain tissues developed at the same pace as the mouse host but maintained rat-like transcriptome profiles. The chimeric rate of rat cells gradually decreased as development progressed, suggesting xenogeneic barriers during mid-to-late pre-natal development. Interspecies forebrain complementation opens the door for studying evolutionarily conserved and divergent mechanisms underlying brain development and cognitive function. The C-CRISPR-based IBC strategy holds great potential to broaden the study and application of interspecies organogenesis.

Mini-dCas13X-mediated RNA editing restores dystrophin expression in a humanized mouse model of Duchenne muscular dystrophy.

Approximately 10% of monogenic diseases are caused by nonsense point mutations that generate premature termination codons (PTCs), resulting in a truncated protein and nonsense-mediated decay of the mutant mRNAs. Here, we demonstrate a mini-dCas13X-mediated RNA adenine base editing (mxABE) strategy to treat nonsense mutation-related monogenic diseases via A-to-G editing in a genetically humanized mouse model of Duchenne muscular dystrophy (DMD). Initially, we identified a nonsense point mutation (c.4174C>T, p.Gln1392*) in the DMD gene of a patient and validated its pathogenicity in humanized mice. In this model, mxABE packaged in a single adeno-associated virus (AAV) reached A-to-G editing rates up to 84% in vivo, at least 20-fold greater than rates reported in previous studies using other RNA editing modalities. Furthermore, mxABE restored robust expression of dystrophin protein to over 50% of WT levels by enabling PTC read-through in multiple muscle tissues. Importantly, systemic delivery of mxABE by AAV also rescued dystrophin expression to averages of 37%, 6%, and 54% of WT levels in the diaphragm, tibialis anterior, and heart muscle, respectively, as well as rescued muscle function. Our data strongly suggest that mxABE-based strategies may be a viable new treatment modality for DMD and other monogenic diseases.

High-fidelity Cas13 variants for targeted RNA degradation with minimal collateral effects.

CRISPR-Cas13 systems have recently been used for targeted RNA degradation in various organisms. However, collateral degradation of bystander RNAs has limited their in vivo applications. Here, we design a dual-fluorescence reporter system for detecting collateral effects and screening Cas13 variants in mammalian cells. Among over 200 engineered variants, several Cas13 variants including Cas13d and Cas13X exhibit efficient on-target activity but markedly reduced collateral activity. Furthermore, transcriptome-wide off-targets and cell growth arrest induced by Cas13 are absent for these variants. High-fidelity Cas13 variants show similar RNA knockdown activity to wild-type Cas13 but no detectable collateral damage in transgenic mice or adeno-associated-virus-mediated somatic cell targeting. Thus, high-fidelity Cas13 variants with minimal collateral effects are now available for targeted degradation of RNAs in basic research and therapeutic applications.

Optimization of C-to-G base editors with sequence context preference predictable by machine learning methods.

Efficient and precise base editors (BEs) for C-to-G transversion are highly desirable. However, the sequence context affecting editing outcome largely remains unclear. Here we report engineered C-to-G BEs of high efficiency and fidelity, with the sequence context predictable via machine-learning methods. By changing the species origin and relative position of uracil-DNA glycosylase and deaminase, together with codon optimization, we obtain optimized C-to-G BEs (OPTI-CGBEs) for efficient C-to-G transversion. The motif preference of OPTI-CGBEs for editing 100 endogenous sites is determined in HEK293T cells. Using a sgRNA library comprising 41,388 sequences, we develop a deep-learning model that accurately predicts the OPTI-CGBE editing outcome for targeted sites with specific sequence context. These OPTI-CGBEs are further shown to be capable of efficient base editing in mouse embryos for generating Tyr-edited offspring. Thus, these engineered CGBEs are useful for efficient and precise base editing, with outcome predictable based on sequence context of targeted sites.

Prediction and Validation of Mouse Meiosis-Essential Genes Based on Spermatogenesis Proteome Dynamics.

The molecular mechanism associated with mammalian meiosis has yet to be fully explored, and one of the main reasons for this lack of exploration is that some meiosis-essential genes are still unknown. The profiling of gene expression during spermatogenesis has been performed in previous studies, yet few studies have aimed to find new functional genes. Since there is a huge gap between the number of genes that are able to be quantified and the number of genes that can be characterized by phenotype screening in one assay, an efficient method to rank quantified genes according to phenotypic relevance is of great importance. We proposed to rank genes by the probability of their function in mammalian meiosis based on global protein abundance using machine learning. Here, nine types of germ cells focusing on continual substages of meiosis prophase I were isolated, and the corresponding proteomes were quantified by high-resolution MS. By combining meiotic labels annotated from the mouse genomics informatics mouse knockout database and the spermatogenesis proteomics dataset, a supervised machine learning package, FuncProFinder (https://github.com/sjq111/FuncProFinder), was developed to rank meiosis-essential candidates. Of the candidates whose functions were unannotated, four of 10 genes with the top prediction scores, Zcwpw1, Tesmin, 1700102P08Rik, and Kctd19, were validated as meiosis-essential genes by knockout mouse models. Therefore, mammalian meiosis-essential genes could be efficiently predicted based on the protein abundance dataset, which provides a paradigm for other functional gene mining from a related abundance dataset.

GOTI, a method to identify genome-wide off-target effects of genome editing in mouse embryos.

Genome editing holds great potential for correcting pathogenic mutations. We developed a method called GOTI (genome-wide off-target analysis by two-cell embryo injection) to detect off-target mutations by editing one blastomere of two-cell mouse embryos using either CRISPR-Cas9 or base editors. GOTI directly compares edited and non-edited cells without the interference of genetic background and thus could detect potential off-target variants with high sensitivity. Notably, the GOTI method was designed to detect potential off-target variants of any genome editing tools by the combination of experimental and computational approaches, which is critical for accurate evaluation of the safety of genome editing tools. Here we provide a detailed protocol for GOTI, including mice mating, two-cell embryo injection, embryonic day 14.5 embryo digestion, fluorescence-activated cell sorting, whole-genome sequencing and data analysis. To enhance the utility of GOTI, we also include a computational workflow called GOTI-seq (https://github.com/sydaileen/GOTI-seq) for the sequencing data analysis, which can generate the final genome-wide off-target variants from raw sequencing data directly. The protocol typically takes 20 d from the mice mating to sequencing and 7 d for sequencing data analysis.

A rationally engineered cytosine base editor retains high on-target activity while reducing both DNA and RNA off-target effects.

Cytosine base editors (CBEs) offer a powerful tool for correcting point mutations, yet their DNA and RNA off-target activities have caused concerns in biomedical applications. We describe screens of 23 rationally engineered CBE variants, which reveal mutation residues in the predicted DNA-binding site can dramatically decrease the Cas9-independent off-target effects. Furthermore, we obtained a CBE variant-YE1-BE3-FNLS-that retains high on-target editing efficiency while causing extremely low off-target edits and bystander edits.

Disruption of splicing-regulatory elements using CRISPR/Cas9 to rescue spinal muscular atrophy in human iPSCs and mice.

We here report a genome-editing strategy to correct spinal muscular atrophy (SMA). Rather than directly targeting the pathogenic exonic mutations, our strategy employed Cas9 and guide-sgRNA for the targeted disruption of intronic splicing-regulatory elements. We disrupted intronic splicing silencers (ISSs, including ISS-N1 and ISS + 100) of survival motor neuron (SMN) 2, a key modifier gene of SMA, to enhance exon 7 inclusion and full-length SMN expression in SMA iPSCs. Survival of splicing-corrected iPSC-derived motor neurons was rescued with SMN restoration. Furthermore, co-injection of Cas9 mRNA from Streptococcus pyogenes (SpCas9) or Cas9 from Staphylococcus aureus (SaCas9) alongside their corresponding sgRNAs targeting ISS-N1 into zygotes rescued 56% and 100% of severe SMA transgenic mice (Smn -/-, SMN2 tg/-). The median survival of the resulting mice was extended to >400 days. Collectively, our study provides proof-of-principle for a new strategy to therapeutically intervene in SMA and other RNA-splicing-related diseases.

Human cleaving embryos enable robust homozygotic nucleotide substitutions by base editors.

Base editing installs a precise nucleotide change in specific gene loci without causing a double-strand break. Its efficiency in human embryos is generally low, limiting its utility in functional genetic studies. Here, we report that injecting base editors into human cleaving two-cell and four-cell embryos results in much higher (up to 13-fold) homozygotic nucleotide substitution efficiency as opposed to MII oocytes or zygotes. Furthermore, as a proof-of-principle study, a point mutation can be efficiently corrected by our method. Our study indicates that human cleaving embryos provide an efficient base editing window for robust gene disruption and correction.

Cytosine base editor generates substantial off-target single-nucleotide variants in mouse embryos.

Genome editing holds promise for correcting pathogenic mutations. However, it is difficult to determine off-target effects of editing due to single-nucleotide polymorphism in individuals. Here we developed a method named GOTI (genome-wide off-target analysis by two-cell embryo injection) to detect off-target mutations by editing one blastomere of two-cell mouse embryos using either CRISPR-Cas9 or base editors. Comparison of the whole-genome sequences of progeny cells of edited and nonedited blastomeres at embryonic day 14.5 showed that off-target single-nucleotide variants (SNVs) were rare in embryos edited by CRISPR-Cas9 or adenine base editor, with a frequency close to the spontaneous mutation rate. By contrast, cytosine base editing induced SNVs at more than 20-fold higher frequencies, requiring a solution to address its fidelity.

Base-Editing-Mediated R17H Substitution in Histone H3 Reveals Methylation-Dependent Regulation of Yap Signaling and Early Mouse Embryo Development.

The coactivator-associated arginine methyltransferase CARM1 catalyzes the methylation of histone H3 arginine 17/26 (H3R17/26me) and non-histone proteins at arginine residues to regulate gene transactivation through profiling or Carm1 overexpression assays. However, the direct relationship between H3R17/26me and its causal role in mouse embryo development remains largely unclear. Here, we use rAPOBEC1-XTEN-Cas9n-UGI (BE3) to efficiently introduce a point mutation (R17H) at multiple Hist1/2H3 loci and a premature-stop codon into the catalytic domain of CARM1 in mouse embryos, resulting in remarkable downregulation of H3R17me levels and developmental defects in pre-implantation and fetal embryos. Transcriptomic analysis reveals that Yap1 and cell cycle signaling pathways are dysregulated in Carm1 truncation and H3R17H substitution embryos, and Yap1 overexpression could rescue the base-editing-elicited defects. Our data establish the direct regulatory relationship between CARM1-mediated H3R17me and early mouse embryo development and demonstrate that Yap1 acts downstream of CARM1-mediated H3R17me to regulate the mouse embryo development.

Simultaneous zygotic inactivation of multiple genes in mouse through CRISPR/Cas9-mediated base editing.

In vivo genetic mutation has become a powerful tool for dissecting gene function; however, multi-gene interaction and the compensatory mechanisms involved can make findings from single mutations, at best difficult to interpret, and, at worst, misleading. Hence, it is necessary to establish an efficient way to disrupt multiple genes simultaneously. CRISPR/Cas9-mediated base editing disrupts gene function by converting a protein-coding sequence into a stop codon; this is referred to as CRISPR-stop. Its application in generating zygotic mutations has not been well explored yet. Here, we first performed a proof-of-principle test by disrupting Atoh1, a gene crucial for auditory hair cell generation. Next, we individually mutated vGlut3 (Slc17a8), otoferlin (Otof) and prestin (Slc26a5), three genes needed for normal hearing function. Finally, we successfully disrupted vGlut3, Otof and prestin simultaneously. Our results show that CRISPR-stop can efficiently generate single or triple homozygous F0 mouse mutants, bypassing laborious mouse breeding. We believe that CRISPR-stop is a powerful method that will pave the way for high-throughput screening of mouse developmental and functional genes, matching the efficiency of methods available for model organisms such as Drosophila.

Tild-CRISPR Allows for Efficient and Precise Gene Knockin in Mouse and Human Cells.

The targeting efficiency of knockin sequences via homologous recombination (HR) is generally low. Here we describe a method we call Tild-CRISPR (targeted integration with linearized dsDNA-CRISPR), a targeting strategy in which a PCR-amplified or precisely enzyme-cut transgene donor with 800-bp homology arms is injected with Cas9 mRNA and single guide RNA into mouse zygotes. Compared with existing targeting strategies, this method achieved much higher knockin efficiency in mouse embryos, as well as brain tissue. Importantly, the Tild-CRISPR method also yielded up to 12-fold higher knockin efficiency than HR-based methods in human embryos, making it suitable for studying gene functions in vivo and developing potential gene therapies.

In vivo simultaneous transcriptional activation of multiple genes in the brain using CRISPR-dCas9-activator transgenic mice.

Despite rapid progresses in the genome-editing field, in vivo simultaneous overexpression of multiple genes remains challenging. We generated a transgenic mouse using an improved dCas9 system that enables simultaneous and precise in vivo transcriptional activation of multiple genes and long noncoding RNAs in the nervous system. As proof of concept, we were able to use targeted activation of endogenous neurogenic genes in these transgenic mice to directly and efficiently convert astrocytes into functional neurons in vivo. This system provides a flexible and rapid screening platform for studying complex gene networks and gain-of-function phenotypes in the mammalian brain.

CRISPR/Cas9-mediated targeted chromosome elimination.

BACKGROUND: The CRISPR/Cas9 system has become an efficient gene editing method for generating cells carrying precise gene mutations, including the rearrangement and deletion of chromosomal segments. However, whether an entire chromosome could be eliminated by this technology is still unknown. RESULTS: Here we demonstrate the use of the CRISPR/Cas9 system to eliminate targeted chromosomes. Using either multiple cleavages induced by a single-guide RNA (sgRNA) that targets multiple chromosome-specific sites or a cocktail of multiple sgRNAs, each targeting one specific site, we found that a sex chromosome could be selectively eliminated in cultured cells, embryos, and tissues in vivo. Furthermore, this approach was able to produce a targeted autosome loss in aneuploid mouse embryonic stem cells with an extra human chromosome and human induced pluripotent stem cells with trisomy 21, as well as cancer cells. CONCLUSIONS: CRISPR/Cas9-mediated targeted chromosome elimination offers a new approach to develop animal models with chromosome deletions, and a potential therapeutic strategy for human aneuploidy diseases involving additional chromosomes.

One-step generation of complete gene knockout mice and monkeys by CRISPR/Cas9-mediated gene editing with multiple sgRNAs.

The CRISPR/Cas9 system is an efficient gene-editing method, but the majority of gene-edited animals showed mosaicism, with editing occurring only in a portion of cells. Here we show that single gene or multiple genes can be completely knocked out in mouse and monkey embryos by zygotic injection of Cas9 mRNA and multiple adjacent single-guide RNAs (spaced 10-200 bp apart) that target only a single key exon of each gene. Phenotypic analysis of F0 mice following targeted deletion of eight genes on the Y chromosome individually demonstrated the robustness of this approach in generating knockout mice. Importantly, this approach delivers complete gene knockout at high efficiencies (100% on Arntl and 91% on Prrt2) in monkey embryos. Finally, we could generate a complete Prrt2 knockout monkey in a single step, demonstrating the usefulness of this approach in rapidly establishing gene-edited monkey models.

Homology-mediated end joining-based targeted integration using CRISPR/Cas9.

Targeted integration of transgenes can be achieved by strategies based on homologous recombination (HR), microhomology-mediated end joining (MMEJ) or non-homologous end joining (NHEJ). The more generally used HR is inefficient for achieving gene integration in animal embryos and tissues, because it occurs only during cell division, although MMEJ and NHEJ can elevate the efficiency in some systems. Here we devise a homology-mediated end joining (HMEJ)-based strategy, using CRISPR/Cas9-mediated cleavage of both transgene donor vector that contains guide RNA target sites and ∼800 bp of homology arms, and the targeted genome. We found no significant improvement of the targeting efficiency by the HMEJ-based method in either mouse embryonic stem cells or the neuroblastoma cell line, N2a, compared to the HR-based method. However, the HMEJ-based method yielded a higher knock-in efficiency in HEK293T cells, primary astrocytes and neurons. More importantly, this approach achieved transgene integration in mouse and monkey embryos, as well as in hepatocytes and neurons in vivo, with an efficiency much greater than HR-, NHEJ- and MMEJ-based strategies. Thus, the HMEJ-based strategy may be useful for a variety of applications, including gene editing to generate animal models and for targeted gene therapies.

CRISPR/Cas9 - Mediated Precise Targeted Integration In Vivo Using a Double Cut Donor with Short Homology Arms.

Precisely targeted genome editing is highly desired for clinical applications. However, the widely used homology-directed repair (HDR)-based genome editing strategies remain inefficient for certain in vivo applications. We here demonstrate a microhomology-mediated end-joining (MMEJ)-based strategy for precisely targeted gene integration in transfected neurons and hepatocytes in vivo with efficiencies up to 20%, much higher (up to 10 fold) than HDR-based strategy in adult mouse tissues. As a proof of concept of its therapeutic potential, we demonstrate the efficacy of MMEJ-based strategy in correction of Fah mutation and rescue of Fah-/- liver failure mice, offering an efficient approach for precisely targeted gene therapies.