Post-translational Regulation of Cas9 during G1 Enhances Homology-Directed Repair

CRISPR RNA-guided autonomous delivery of Cas9

Cas9 is an endonuclease that can be programed to autonomously deliver diverse effectors to specified genetic addresses. High-resolution structures of this protein and its associated CRISPR RNA guide explain the molecular mechanisms of CRISPR-RNA-guided DNA recognition and provide a molecular blueprint that has facilitated structure-guided functional remodeling. Here we retrace events that led from early efforts to understand the central role of Cas9 in CRISPR-mediated adaptive immunity to contemporary efforts aimed at developing and deploying this enzyme for programmable genetic editing.

The road less traveled: strategies to enhance the frequency of homology-directed repair (HDR) for increased efficiency of CRISPR/Cas-mediated transgenesis

Non-homologous end joining (NHEJ), and to a lesser extent, the error-free pathway known as homology-directed repair (HDR) are cellular mechanisms for recovery from double-strand DNA breaks (DSB) induced by RNA-guided programmable nuclease CRISPR/Cas. Since NHEJ is equivalent to using a duck tape to stick two pieces of metals together, the outcome of this repair mechanism is prone to error. Any out-of-frame mutations or premature stop codons resulting from NHEJ repair mechanism are extremely handy for loss-of-function studies. Substitution of a mutation on the genome with the correct exogenous repair DNA requires coordination via an error-free HDR, for targeted transgenesis. However, several practical limitations exist in harnessing the potential of HDR to replace a faulty mutation for therapeutic purposes in all cell types and more so in somatic cells. In germ cells after the DSB, copying occurs from the homologous chromosome, which increases the chances of incorporation of exogenous DNA with some degree of homology into the genome compared with somatic cells where copying from the identical sister chromatid is always preferred. This review summarizes several strategies that have been implemented to increase the frequency of HDR with a focus on somatic cells. It also highlights the limitations of this technology in gene therapy and suggests specific solutions to circumvent those barriers.

Transient Retrovirus-Based CRISPR/Cas9 All-in-One Particles for Efficient, Targeted Gene Knockout

The recently discovered CRISPR/Cas9 system is widely used in basic research and is a useful tool for disease modeling and gene editing therapies. However, long-term expression of DNA-modifying enzymes can be associated with cytotoxicity and is particularly unwanted in clinical gene editing strategies. Because current transient expression methods may still suffer from cytotoxicity and/or low efficiency, we developed non-integrating retrovirus-based CRISPR/Cas9 all-in-one particles for targeted gene knockout. By redirecting the gammaretroviral packaging machinery, we transiently delivered Streptococcus pyogenes Cas9 (SpCas9) mRNA and single-guide RNA transcripts into various (including primary) cell types. Spatiotemporal co-delivery of CRISPR/Cas9 components resulted in efficient disruption of a surrogate reporter gene, as well as functional knockout of endogenous human genes CXCR4 and TP53. Although acting in a hit-and-run fashion, knockout efficiencies of our transient particles corresponded to 52%-80% of those obtained from constitutively active integrating vectors. Stable SpCas9 overexpression at high doses in murine NIH3T3 cells caused a substantial G0/G1 arrest accompanied by reduced cell growth and metabolic activity, which was prevented by transient SpCas9 transfer. In summary, the non-integrating retrovirus-based vector particles introduced here allow efficient and dose-controlled delivery of CRISPR/Cas9 components into target cells.

Shortening the Half-Life of Cas9 Maintains Its Gene Editing Ability and Reduces Neuronal Toxicity

Virus-mediated expression of CRISPR/Cas9 is commonly used for genome editing in animal brains to model or treat neurological diseases, but the potential neurotoxicity of overexpressing bacterial Cas9 in the mammalian brain remains unknown. Through RNA sequencing (RNA-seq) analysis, we find that virus-mediated expression of Cas9 influences the expression of genes involved in neuronal functions. Reducing the half-life of Cas9 by tagging with geminin, whose expression is regulated by the cell cycle, maintains the genome editing capacity of Cas9 but significantly alleviates neurotoxicity. Thus, modification of Cas9 by shortening its half-life can help develop CRISPR/Cas9-based therapeutic approaches for treating neurological disorders.

CRISPR and Target-Specific DNA Endonucleases for Efficient DNA Knock-in in Eukaryotic Genomes

The discovery and mechanistic understanding of target-specific genome engineering technologies has led to extremely effective and specific genome editing in higher organisms. Target-specific genetic modification technology is expected to have a leading position in future gene therapy development, and has a ripple effect on various basic and applied studies. However, several problems remain and hinder efficient and specific editing of target genomic loci. The issues are particularly critical in precise targeted insertion of external DNA sequences into genomes. Here, we discuss some recent efforts to overcome such problems and present a perspective of future genome editing technologies.

Improving Gene Editing Outcomes in Human Hematopoietic Stem and Progenitor Cells by Temporal Control of DNA Repair

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated system (Cas9)-mediated gene editing of human hematopoietic stem cells (hHSCs) is a promising strategy for the treatment of genetic blood diseases through site-specific correction of identified causal mutations. However, clinical translation is hindered by low ratio of precise gene modification using the corrective donor template (homology-directed repair, HDR) to gene disruption (nonhomologous end joining, NHEJ) in hHSCs. By using a modified version of Cas9 with reduced nuclease activity in G1 phase of cell cycle when HDR cannot occur, and transiently increasing the proportion of cells in HDR-preferred phases (S/G2), we achieved a four-fold improvement in HDR/NHEJ ratio over the control condition in vitro, and a significant improvement after xenotransplantation of edited hHSCs into immunodeficient mice. This strategy for improving gene editing outcomes in hHSCs has important implications for the field of gene therapy, and can be applied to diseases where increased HDR/NHEJ ratio is critical for therapeutic success. Stem Cells 2019;37:284-294.

Critical Role of the UBL Domain in Stimulating the E3 Ubiquitin Ligase Activity of UHRF1 toward Chromatin

The RING E3 ubiquitin ligase UHRF1 controls DNA methylation through its ability to target the maintenance DNA methyltransferase DNMT1 to newly replicated chromatin. DNMT1 recruitment relies on ubiquitylation of histone H3 by UHRF1; however, how UHRF1 deposits ubiquitin onto the histone is unknown. Here, we demonstrate that the ubiquitin-like domain (UBL) of UHRF1 is essential for RING-mediated H3 ubiquitylation. Using chemical crosslinking and mass spectrometry, biochemical assays, and recombinant chromatin substrates, we show that the UBL participates in structural rearrangements of UHRF1 upon binding to chromatin and the E2 ubiquitin conjugating enzyme UbcH5a/UBE2D1. Similar to ubiquitin, the UBL exerts its effects through a hydrophobic patch that contacts a regulatory surface on the “backside” of the E2 to stabilize the E2-E3-chromatin complex. Our analysis of the enzymatic mechanism of UHRF1 uncovers an unexpected function of the UBL domain and defines a new role for this domain in DNMT1-dependent inheritance of DNA methylation.

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The UBL domain of UHRF1 is required for its E3 ubiquitin ligase activity

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A hydrophobic patch on the UBL is required to form a stable E2/E3/chromatin complex

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The UHRF1 N terminus and UBL hydrophobic patch control targeted H3 ubiquitylation

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DNMT1-mediated maintenance DNA methylation requires the UBL hydrophobic patch

Highly efficient genome editing by homology-directed repair using Cas9 protein in Ceratitis capitata

The Mediterranean fruit fly Ceratitis capitata is a highly polyphagous and invasive insect pest, causing enormous economic damage in horticultural systems. A successful and environment-friendly control strategy is the sterile insect technique (SIT) that reduces pest populations through infertile matings with mass-released, sterilized insects. However, the SIT is not readily applicable to each pest species. While transgenic approaches hold great promise to improve critical aspects of the SIT to transfer it to new species, they are suspect to strict or even prohibitive legislation regarding the release of genetically modified (GM) organisms. In contrast, specific mutations created via CRISPR-Cas genome editing are not regulated as GM in the US, and might thus allow creating optimal strains for SIT. Here, we describe highly efficient homology-directed repair genome editing in C. capitata by injecting pre-assembled CRISPR-Cas9 ribonucleoprotein complexes using different guide RNAs and a short single-stranded oligodeoxynucleotide donor to convert an enhanced green fluorescent protein in C. capitata into a blue fluorescent protein. Six out of seven fertile and individually backcrossed G0 individuals generated 57-90% knock-in rate within their total offspring and 70-96% knock-in rate within their phenotypically mutant offspring. Based on the achieved efficiency, this approach could also be used to introduce mutations which do not produce a screenable phenotype and identify positive mutants with a reasonable workload. Furthermore, CRISPR-Cas HDR would allow to recreate mutations formerly identified in classical mutagenesis screens and to transfer them to related species to establish new (SIT-like) pest control systems. Considering the potential that CRISPR-induced alterations in organisms could be classified as non-GM in additional countries, such new strains could potentially be used for pest control applications without the need to struggle with GMO directives.

Decoding non-random mutational signatures at Cas9 targeted sites

The mutation patterns at Cas9 targeted sites contain unique information regarding the nuclease activity and repair mechanisms in mammalian cells. However, analytical framework for extracting such information are lacking. Here, we present a novel computational platform called Rational InDel Meta-Analysis (RIMA) that enables an in-depth comprehensive analysis of Cas9-induced genetic alterations, especially InDels mutations. RIMA can be used to quantitate the contribution of classical microhomology-mediated end joining (c-MMEJ) pathway in the formation of mutations at Cas9 target sites. We used RIMA to compare mutational signatures at 15 independent Cas9 target sites in human A549 wildtype and A549-POLQ knockout cells to elucidate the role of DNA polymerase θ in c-MMEJ. Moreover, the single nucleotide insertions at the Cas9 target sites represent duplications of preceding nucleotides, suggesting that the flexibility of the Cas9 nuclease domains results in both blunt- and staggered-end cuts. Thymine at the fourth nucleotide before protospacer adjacent motif (PAM) results in a two-fold higher occurrence of single nucleotide InDels compared to guanine at the same position. This study provides a novel approach for the characterization of the Cas9 nucleases with improved accuracy in predicting genome editing outcomes and a potential strategy for homology-independent targeted genomic integration.

The road less traveled: strategies to enhance the frequency of homology-directed repair (HDR) for increased efficiency of CRISPR/Cas-mediated transgenesis

Non-homologous end joining (NHEJ), and to a lesser extent, the error-free pathway known as homology-directed repair (HDR) are cellular mechanisms for recovery from double-strand DNA breaks (DSB) induced by RNA-guided programmable nuclease CRISPR/Cas. Since NHEJ is equivalent to using a duck tape to stick two pieces of metals together, the outcome of this repair mechanism is prone to error. Any out-of-frame mutations or premature stop codons resulting from NHEJ repair mechanism are extremely handy for loss-of-function studies. Substitution of a mutation on the genome with the correct exogenous repair DNA requires coordination via an error-free HDR, for targeted transgenesis. However, several practical limitations exist in harnessing the potential of HDR to replace a faulty mutation for therapeutic purposes in all cell types and more so in somatic cells. In germ cells after the DSB, copying occurs from the homologous chromosome, which increases the chances of incorporation of exogenous DNA with some degree of homology into the genome compared with somatic cells where copying from the identical sister chromatid is always preferred. This review summarizes several strategies that have been implemented to increase the frequency of HDR with a focus on somatic cells. It also highlights the limitations of this technology in gene therapy and suggests specific solutions to circumvent those barriers. [BMB Reports 2018; 51(9): 437-443].

Increasing the precision of gene editing in vitro, ex vivo, and in vivo

New gene editing tools like CRISPR-Cas9 enable precision genome engineering within cell lines, primary cells, and model organisms, with some formulations now entering the clinic. "Precision" applies to various aspects of gene editing, and can be tailored for each application. Here we review recent advances in four types of precision in gene editing: 1) increased DNA cutting precision (e.g., on-target:off-target nuclease specificity), 2) increased on-target knock-in of sequence variants and transgenes (e.g., increased homology-directed repair), 3) increased transcriptional control of edited genes, and 4) increased specificity in delivery to a specific cell or tissue. Design of next-generation gene and cell therapies will likely exploit a combination of these advances.

Efficient single-copy HDR by 5' modified long dsDNA donors

CRISPR/Cas9 efficiently induces targeted mutations via non-homologous-end-joining but for genome editing, precise, homology-directed repair (HDR) of endogenous DNA stretches is a prerequisite. To favor HDR, many approaches interfere with the repair machinery or manipulate Cas9 itself. Using Medaka we show that the modification of 5’ ends of long dsDNA donors strongly enhances HDR, favors efficient single-copy integration by retaining a monomeric donor conformation thus facilitating successful gene replacement or tagging.

CRISPR/Cas9 technology has revolutionized the ability of researchers to edit the DNA of any organism whose genome has already been sequenced. In the editing process, a section of RNA acts as a guide to match up to the location of the target DNA. The enzyme Cas9 then makes a cut in both strands of the DNA at this specific location. New segments of DNA can be introduced to the cell, incorporated into DNA ‘templates’. The cell uses the template to help it to heal the double-strand break, and in doing so adds the new DNA segment into the organism’s genome.

A drawback of CRISPR/Cas9 is that it often introduces multiple copies of the new DNA segment into the genome because the templates can bind to each other before being pasted into place. In addition, some parts of the new DNA segment can be missed off during the editing process. However, most applications of CRISPR/Cas9 – for example, to replace a defective gene with a working version – require exactly one whole copy of the desired DNA to be inserted into the genome.

In order to achieve more accurate CRISPR/Cas9 genome editing, Gutierrez-Triana, Tavhelidse, Thumberger et al. attached additional molecules to the end of the DNA template to shield the DNA from mistakes during editing. The modified template was used to couple a stem cell gene to a reporter that produces a green fluorescent protein into the genome of fish embryos. The fluorescent proteins made it easy to identify when the coupling was successful.

Gutierrez-Triana et al. found that the additional molecules prevented multiple templates from joining together end to end, and ensured the full DNA segment was inserted into the genome. Furthermore, the results of the experiments showed that only one copy of the template was inserted into the DNA of the fish. In the future, the new template will allow DNA to be edited in a more controlled way both in basic research and in therapeutic applications.

Programmable DNA repair with CRISPRa/i enhanced homology-directed repair efficiency with a single Cas9

CRISPR systems have been proven as versatile tools for site-specific genome engineering in mammalian species. During the gene editing processes, these RNA-guide nucleases introduce DNA double strand breaks (DSBs), in which non-homologous DNA end joining (NHEJ) dominates the DNA repair pathway, limiting the efficiency of homology-directed repair (HDR), the alternative pathway essential for precise gene targeting. Multiple approaches have been developed to enhance HDR, including chemical compound or RNA interference-mediated inhibition of NHEJ factors, small molecule activation of HDR enzymes, or cell cycle timed delivery of CRISPR complex. However, these approaches face multiple challenges, yet have moderate or variable effects. Here we developed a new approach that programs both NHEJ and HDR pathways with CRISPR activation and interference (CRISPRa/i) to achieve significantly enhanced HDR efficiency of CRISPR-mediated gene editing. The manipulation of NHEJ and HDR pathway components, such as CtIP, CDK1, KU70, KU80, and LIG4, was mediated by catalytically dead guide RNAs (dgRNAs), thus relying on only a single catalytically active Cas9 to perform both CRISPRa/i and precise gene editing. While reprogramming of most DNA repair factors or their combinations tested enhanced HDR efficiency, simultaneously activating CDK1 and repressing KU80 has the strongest effect with increased HDR rate upto an order of magnitude. Doxycycline-induced dgRNA-based CRISPRa/i programming of DNA repair enzymes, as well as viral packaging enabled flexible and tunable HDR enhancement for broader applicability in mammalian cells. Our study provides an effective, flexible, and potentially safer strategy to enhance precise genome modifications, which might broadly impact human gene editing and therapy.

Transgenic Mouse Models in Cancer Research

The use of existing mouse models in cancer research is of utmost importance as they aim to explore the casual link between candidate cancer genes and carcinogenesis as well as to provide models to develop and test new therapies. However, faster progress in translating mouse cancer model research into the clinic has been hampered due to the limitations of these models to better reflect the complexities of human tumors. Traditionally, immunocompetent and immunodeficient mice with syngeneic and xenografted tumors transplanted subcutaneously or orthotopically have been used. These models are still being widely employed for many different types of studies, in part due to their widespread availability and low cost. Other types of mouse models used in cancer research comprise transgenic mice in which oncogenes can be constitutively or conditionally expressed and tumor-suppressor genes silenced using conventional methods, such as retroviral infection, microinjection of DNA constructs, and the so-called "gene-targeted transgene" approach. These traditional transgenic models have been very important in studies of carcinogenesis and tumor pathogenesis, as well as in studies evaluating the development of resistance to therapy. Recently, the clustered regularly interspaced short palindromic repeats (CRISPR)-based genome editing approach has revolutionized the field of mouse cancer models and has had a profound and rapid impact on the development of more effective systems to study human cancers. The CRISPR/Cas9-based transgenic models have the capacity to engineer a wide spectrum of mutations found in human cancers and provide solutions to problems that were previously unsolvable. Recently, humanized mouse xenograft models that accept patient-derived xenografts and CD34+ cells were developed to better mimic tumor heterogeneity, the tumor microenvironment, and cross-talk between the tumor and stromal/immune cells. These features make them extremely valuable models for the evaluation of investigational cancer therapies, specifically new immunotherapies. Taken together, improvements in both the CRISPR/Cas9 system producing more valid mouse models and in the humanized mouse xenograft models resembling complex interactions between the tumor and its environment might represent one of the successful pathways to precise individualized cancer therapy, leading to improved cancer patient survival and quality of life.

Genome editing by natural and engineered CRISPR-associated nucleases

Over the last decade, research on distinct types of CRISPR systems has revealed many structural and functional variations. Recently, several novel types of single-polypeptide CRISPR-associated systems have been discovered including Cas12a/Cpf1 and Cas13a/C2c2. Despite distant similarities to Cas9, these additional systems have unique structural and functional features, providing new opportunities for genome editing applications. Here, relevant fundamental features of natural and engineered CRISPR-Cas variants are compared. Moreover, practical matters are discussed that are essential for dedicated genome editing applications, including nuclease regulation and delivery, target specificity, as well as host repair diversity.

Efficient Knock-in of a Point Mutation in Porcine Fibroblasts Using the CRISPR/Cas9-GMNN Fusion Gene

During CRISPR/Cas9 mediated genome editing, site-specific double strand breaks are introduced and repaired either unspecific by non-homologous end joining (NHEJ) or sequence dependent by homology directed repair (HDR). Whereas NHEJ-based generation of gene knock-out is widely performed, the HDR-based knock-in of specific mutations remains a bottleneck. Especially in primary cell lines that are essential for the generation of cell culture and animal models of inherited human diseases, knock-in efficacy is insufficient and needs significant improvement. Here, we tested two different approaches to increase the knock-in frequency of a specific point mutation into the MYH7-gene in porcine fetal fibroblasts. We added a small molecule inhibitor of NHEJ, SCR7 (5,6-bis((E)-benzylideneamino)-2-mercaptopyrimidin-4-ol), during genome editing and screened cell cultures for the point mutation. However, this approach did not yield increased knock-in rates. In an alternative approach, we fused humanized Cas9 (hCas9) to the N-terminal peptide of the Geminin gene (GMNN). The fusion protein is degraded in NHEJ-dominated cell cycle phases, which should increase HDR-rates. Using hCas9-GMNN and point mutation-specific real time PCR screening, we found a two-fold increase in genome edited cell cultures. This increase of HDR by hCas9-GMNN provides a promising way to enrich specific knock-in in porcine fibroblast cultures for somatic cloning approaches.

Increasing Cas9-mediated homology-directed repair efficiency through covalent tethering of DNA repair template

The CRISPR-Cas9 system is a powerful genome-editing tool in which a guide RNA targets Cas9 to a site in the genome, where the Cas9 nuclease then induces a double-stranded break (DSB). The potential of CRISPR-Cas9 to deliver precise genome editing is hindered by the low efficiency of homology-directed repair (HDR), which is required to incorporate a donor DNA template encoding desired genome edits near the DSB. We present a strategy to enhance HDR efficiency by covalently tethering a single-stranded oligodeoxynucleotide (ssODN) to the Cas9-guide RNA ribonucleoprotein (RNP) complex via a fused HUH endonuclease, thus spatially and temporally co-localizing the DSB machinery and donor DNA. We demonstrate up to a 30-fold enhancement of HDR using several editing assays, including repair of a frameshift and in-frame insertions of protein tags. The improved HDR efficiency is observed in multiple cell types and target loci and is more pronounced at low RNP concentrations.

Systematic evaluation of CRISPR-Cas systems reveals design principles for genome editing in human cells

BACKGROUND

While CRISPR-Cas systems hold tremendous potential for engineering the human genome, it is unclear how well each system performs against one another in both non-homologous end joining (NHEJ)-mediated and homology-directed repair (HDR)-mediated genome editing.

RESULTS

We systematically compare five different CRISPR-Cas systems in human cells by targeting 90 sites in genes with varying expression levels. For a fair comparison, we select sites that are either perfectly matched or have overlapping seed regions for Cas9 and Cpf1. Besides observing a trade-off between cleavage efficiency and target specificity for these natural endonucleases, we find that the editing activities of the smaller Cas9 enzymes from Staphylococcus aureus (SaCas9) and Neisseria meningitidis (NmCas9) are less affected by gene expression than the other larger Cas proteins. Notably, the Cpf1 nucleases from Acidaminococcus sp. BV3L6 and Lachnospiraceae bacterium ND2006 (AsCpf1 and LbCpf1, respectively) are able to perform precise gene targeting efficiently across multiple genomic loci using single-stranded oligodeoxynucleotide (ssODN) donor templates with homology arms as short as 17 nucleotides. Strikingly, the two Cpf1 nucleases exhibit a preference for ssODNs of the non-target strand sequence, while the popular Cas9 enzyme from Streptococcus pyogenes (SpCas9) exhibits a preference for ssODNs of the target strand sequence instead. Additionally, we find that the HDR efficiencies of Cpf1 and SpCas9 can be further improved by using asymmetric donors with longer arms 5' of the desired DNA changes.

CONCLUSIONS

Our work delineates design parameters for each CRISPR-Cas system and will serve as a useful reference for future genome engineering studies.

Covalent linkage of the DNA repair template to the CRISPR-Cas9 nuclease enhances homology-directed repair

The CRISPR-Cas9 targeted nuclease technology allows the insertion of genetic modifications with single base-pair precision. The preference of mammalian cells to repair Cas9-induced DNA double-strand breaks via error-prone end-joining pathways rather than via homology-directed repair mechanisms, however, leads to relatively low rates of precise editing from donor DNA. Here we show that spatial and temporal co-localization of the donor template and Cas9 via covalent linkage increases the correction rates up to 24-fold, and demonstrate that the effect is mainly caused by an increase of donor template concentration in the nucleus. Enhanced correction rates were observed in multiple cell types and on different genomic loci, suggesting that covalently linking the donor template to the Cas9 complex provides advantages for clinical applications where high-fidelity repair is desired.

Applying the auxin-inducible degradation system for rapid protein depletion in mammalian cells

The ability to deplete a protein of interest is critical for dissecting cellular processes. Traditional methods of protein depletion are often slow acting, which can be problematic when characterizing a cellular process that occurs within a short period of time, such as mitosis. Furthermore, these methods are usually not reversible. Recent advances to achieve protein depletion function by inducibly trafficking proteins of interest to an endogenous E3 ubiquitin ligase complex to promote ubiquitination and subsequent degradation by the proteasome. One of these systems, the auxin-inducible degron (AID) system, has been shown to permit rapid and inducible degradation of AID-tagged target proteins in mammalian cells. The AID system can control the abundance of a diverse set of cellular proteins, including those contained within protein complexes, and is active in all phases of the cell cycle. Here we discuss considerations for the successful implementation of the AID system and describe a protocol using CRISPR/Cas9 to achieve biallelic insertion of an AID in human cells. This method can also be adapted to insert other tags, such as fluorescent proteins, at defined genomic locations.

CtIP fusion to Cas9 enhances transgene integration by homology-dependent repair

In genome editing with CRISPR-Cas9, transgene integration often remains challenging. Here, we present an approach for increasing the efficiency of transgene integration by homology-dependent repair (HDR). CtIP, a key protein in early steps of homologous recombination, is fused to Cas9 and stimulates transgene integration by HDR at the human AAVS1 safe harbor locus. A minimal N-terminal fragment of CtIP, designated HE for HDR enhancer, is sufficient to stimulate HDR and this depends on CDK phosphorylation sites and the multimerization domain essential for CtIP activity in homologous recombination. HDR stimulation by Cas9-HE, however, depends on the guide RNA used, a limitation that may be overcome by testing multiple guides to the locus of interest. The Cas9-HE fusion is simple to use and allows obtaining twofold or more efficient transgene integration than that with Cas9 in several experimental systems, including human cell lines, iPS cells, and rat zygotes.

An efficient platform for generating somatic point mutations with germline transmission in the zebrafish by CRISPR/Cas9-mediated gene editing

Homology-directed recombination (HDR)-mediated genome editing is a powerful approach for both basic functional study and disease modeling. Although some studies have reported HDR-mediated precise editing in nonrodent models, the efficiency of establishing pure mutant animal lines that carry specific amino acid substitutions remains low. Furthermore, because the efficiency of nonhomologous end joining (NHEJ)-induced insertion and deletion (indel) mutations is normally much higher than that of HDR-induced point mutations, it is often difficult to identify the latter in the background of indel mutations. Using zebrafish as the model organism and Y box-binding protein 1 (Ybx1/ybx1) as the model molecule, we have established an efficient platform for precise CRISPR/Cas9-mediated gene editing in somatic cells, yielding an efficiency of up to 74% embryos. Moreover, we established a procedure for screening germline transmission of point mutations out of indel mutations even when germline transmission efficiency was low (<2%). To further improve germline transmission of HDR-induced point mutations, we optimized several key factors that may affect HDR efficiency, including the type of DNA donor, suppression of NHEJ, stimulation of HDR pathways, and use of Cas9 protein instead of mRNA. The optimized combination of these factors significantly increased the efficiency of germline transmission of point mutation up to 25%. In summary, we have developed an efficient procedure for creating point mutations and differentiating mutant individuals from those carrying knockouts of entire genes.

Genome Editing: Insights from Chemical Biology to Support Safe and Transformative Therapeutic Applications

Programmable nuclease-based genome editing technologies, including the clustered, regularly interspaced, short palindromic repeats (CRISPR)/Cas9 system, are becoming an essential component of many applications ranging from agriculture to medicine. However, fundamental limitations currently prevent the widespread, safe, and practical use of genome editors, especially for human disease interventions. These limitations include off-target effects, a lack of control over editing activity, suboptimal DNA repair outcomes, insufficient target conversion, and inadequate delivery performance. This perspective focuses on the potential for biological chemistry to address these limitations such that newly developed genome editing technologies can enable the broadest range of potential future applications. Equally important will be the development of these powerful technologies within a relevant ethical framework that emphasizes safety and responsible innovation.

Modulating DNA Repair Pathways to Improve Precision Genome Engineering

Programmable nucleases like the popular CRISPR/Cas9 system allow for precision genome engineering by inducing a site-specific DNA double strand break (DSB) within a genome. The DSB is repaired by endogenous DNA repair pathways, either nonhomologous end joining (NHEJ) or homology directed repair (HDR). The predominant and error-prone NHEJ pathway often results in small nucleotide insertions or deletions that can be used to construct knockout alleles. Alternatively, HDR activity can result in precise modification incorporating exogenous DNA fragments into the cut site. However, genetic recombination in mammalian systems through the HDR pathway is an inefficient process and requires cumbersome laboratory methods to identify the desired accurate insertion events. This is further compromised by the activity of the competing DNA repair pathway, NHEJ, which repairs the majority of nuclease induced DNA DSBs and also is responsible for mutagenic insertion and deletion events at off-target locations throughout the genome. Various methodologies have been developed to increase the efficiency of designer nuclease-based HDR mediated gene editing. Here, we review these advances toward modulating the activities of the two critical DNA repair pathways, HDR and NHEJ, to enhance precision genome engineering.

Use of CRISPR/Cas9 to model brain diseases

Aging-related brain diseases consist of a number of important neurodegenerative disorders, including Alzheimer's, Parkinson's, and Huntington's diseases, all of which have become more prevalent as the life expectancy of humans is prolonged. Age-dependent brain disorders are associated with both environmental insults and genetic mutations. For those brain disorders that are inherited, gene editing is an important tool for establishing animal models to investigate the pathogenesis of disease and identify effective treatments. Here we focus on the tools for gene editing, especially CRISPR/Cas9, and discuss their application for generating animal models that can recapitulate the brain pathology seen in human diseases. We also highlight the advantages and disadvantages of establishing genetically modified animal models. Finally, we discuss recent findings to resolve technical issues related to the use of CRISPR/Cas9 for generating animal models of brain diseases.

CRISPR-engineered genome editing for the next generation neurological disease modeling

Neurological disorders often occur because of failure of proper brain development and/or appropriate maintenance of neuronal circuits. In order to understand roles of causative factors (e.g. genes, cell types) in disease development, generation of solid animal models has been one of straight-forward approaches. Recent next generation sequencing studies on human patient-derived clinical samples have identified various types of recurrent mutations in individual neurological diseases. While these discoveries have prompted us to evaluate impact of mutated genes on these neurological diseases, a feasible but flexible genome editing tool had remained to be developed. An advance of genome editing technology using the clustered regularly interspaced short palindromic repeats (CRISPR) with the CRISPR-associated protein (Cas) offers us a tremendous potential to create a variety of mutations in the cell, leading to "next generation" disease models carrying disease-associated mutations. We will here review recent progress of CRISPR-based brain disease modeling studies and discuss future requirement to tackle current difficulties in usage of these technologies.

Applications of Alternative Nucleases in the Age of CRISPR/Cas9

Breakthroughs in the development of programmable site-specific nucleases, including zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), meganucleases (MNs), and most recently, the clustered regularly interspaced short palindromic repeats (CRISPR) associated proteins (including Cas9) have greatly enabled and accelerated genome editing. By targeting double-strand breaks to user-defined locations, the rates of DNA repair events are greatly enhanced relative to un-catalyzed events at the same sites. However, the underlying biology of each genome-editing nuclease influences the targeting potential, the spectrum of off-target cleavages, the ease-of-use, and the types of recombination events at targeted double-strand breaks. No single genome-editing nuclease is optimized for all possible applications. Here, we focus on the diversity of nuclease domains available for genome editing, highlighting biochemical properties and the potential applications that are best suited to each domain.

Disruption of diphthamide synthesis genes and resulting toxin resistance as a robust technology for quantifying and optimizing CRISPR/Cas9-mediated gene editing

We have devised an effective and robust method for the characterization of gene-editing events. The efficacy of editing-mediated mono- and bi-allelic gene inactivation and integration events is quantified based on colony counts. The combination of diphtheria toxin (DT) and puromycin (PM) selection enables analyses of 10,000-100,000 individual cells, assessing hundreds of clones with inactivated genes per experiment. Mono- and bi-allelic gene inactivation is differentiated by DT resistance, which occurs only upon bi-allelic inactivation. PM resistance indicates integration. The robustness and generalizability of the method were demonstrated by quantifying the frequency of gene inactivation and cassette integration under different editing approaches: CRISPR/Cas9-mediated complete inactivation was ~30-50-fold more frequent than cassette integration. Mono-allelic inactivation without integration occurred >100-fold more frequently than integration. Assessment of gRNA length confirmed 20mers to be most effective length for inactivation, while 16-18mers provided the highest overall integration efficacy. The overall efficacy was ~2-fold higher for CRISPR/Cas9 than for zinc-finger nuclease and was significantly increased upon modulation of non-homologous end joining or homology-directed repair. The frequencies and ratios of editing events were similar for two different DPH genes (independent of the target sequence or chromosomal location), which indicates that the optimization parameters identified with this method can be generalized.

Hematopoietic Stem Cell Gene Therapy: Progress and Lessons Learned

The use of allogeneic hematopoietic stem cells (HSCs) to treat genetic blood cell diseases has become a clinical standard but is limited by the availability of suitable matched donors and potential immunologic complications. Gene therapy using autologous HSCs should avoid these limitations and thus may be safer. Progressive improvements in techniques for genetic correction of HSCs, by either vector gene addition or gene editing, are facilitating successful treatments for an increasing number of diseases. We highlight the progress, successes, and remaining challenges toward the development of HSC gene therapies and discuss lessons they provide for the development of future clinical stem cell therapies.

Targeted gene knock-in by homology-directed genome editing using Cas9 ribonucleoprotein and AAV donor delivery

Realizing the full potential of genome editing requires the development of efficient and broadly applicable methods for delivering programmable nucleases and donor templates for homology-directed repair (HDR). The RNA-guided Cas9 endonuclease can be introduced into cells as a purified protein in complex with a single guide RNA (sgRNA). Such ribonucleoproteins (RNPs) can facilitate the high-fidelity introduction of single-base substitutions via HDR following co-delivery with a single-stranded DNA oligonucleotide. However, combining RNPs with transgene-containing donor templates for targeted gene addition has proven challenging, which in turn has limited the capabilities of the RNP-mediated genome editing toolbox. Here, we demonstrate that combining RNP delivery with naturally recombinogenic adeno-associated virus (AAV) donor vectors enables site-specific gene insertion by homology-directed genome editing. Compared to conventional plasmid-based expression vectors and donor templates, we show that combining RNP and AAV donor delivery increases the efficiency of gene addition by up to 12-fold, enabling the creation of lineage reporters that can be used to track the conversion of striatal neurons from human fibroblasts in real time. These results thus illustrate the potential for unifying nuclease protein delivery with AAV donor vectors for homology-directed genome editing.

Updated summary of genome editing technology in human cultured cells linked to human genetics studies

Current deep-sequencing technology provides a mass of nucleotide variations associated with human genetic disorders to accelerate the identification of causative mutations. To understand the etiology of genetic disorders, reverse genetics in human cultured cells is a useful approach for modeling a disease in vitro. However, gene targeting in human cultured cells is difficult because of their low activity of homologous recombination. Engineered endonucleases enable enhancement of the local activation of DNA repair pathways at the human genome target site to rewrite the desired sequence, thereby efficiently generating disease-modeling cultured cell clones. These edited cells can be used to explore the molecular functions of a causative gene product to uncover the etiological mechanisms. The correction of mutations in patient cells using genome editing technology could contribute to the development of unique gene therapies. This technology can also be applied to screening causative mutations. Rare genetic disorders and non-exonic mutation-caused diseases remain frontier in the field of human genetics as it is difficult to validate whether the extracted nucleotide variants are mutation or polymorphism. When isogenic human cultured cells with a candidate variant reproduce the pathogenic phenotypes, it is confirmed that the variant is a causative mutation.

Engineered CRISPR Systems for Next Generation Gene Therapies

An ideal in vivo gene therapy platform provides safe, reprogrammable, and precise strategies which modulate cell and tissue gene regulatory networks with a high temporal and spatial resolution. Clustered regularly interspaced short palindromic repeats (CRISPR), a bacterial adoptive immune system, and its CRISPR-associated protein 9 (Cas9), have gained attention for the ability to target and modify DNA sequences on demand with unprecedented flexibility and precision. The precision and programmability of Cas9 is derived from its complexation with a guide-RNA (gRNA) that is complementary to a desired genomic sequence. CRISPR systems open-up widespread applications including genetic disease modeling, functional screens, and synthetic gene regulation. The plausibility of in vivo genetic engineering using CRISPR has garnered significant traction as a next generation in vivo therapeutic. However, there are hurdles that need to be addressed before CRISPR-based strategies are fully implemented. Some key issues center on the controllability of the CRISPR platform, including minimizing genomic-off target effects and maximizing in vivo gene editing efficiency, in vivo cellular delivery, and spatial-temporal regulation. The modifiable components of CRISPR systems: Cas9 protein, gRNA, delivery platform, and the form of CRISPR system delivered (DNA, RNA, or ribonucleoprotein) have recently been engineered independently to design a better genome engineering toolbox. This review focuses on evaluating CRISPR potential as a next generation in vivo gene therapy platform and discusses bioengineering advancements that can address challenges associated with clinical translation of this emerging technology.

Towards mastering CRISPR-induced gene knock-in in plants: Survey of key features and focus on the model Physcomitrella patens

Beyond its predominant role in human and animal therapy, the CRISPR-Cas9 system has also become an essential tool for plant research and plant breeding. Agronomic applications rely on the mastery of gene inactivation and gene modification. However, if the knock-out of genes by non-homologous end-joining (NHEJ)-mediated repair of the targeted double-strand breaks (DSBs) induced by the CRISPR-Cas9 system is rather well mastered, the knock-in of genes by homology-driven repair or end-joining remains difficult to perform efficiently in higher plants. In this review, we describe the different approaches that can be tested to improve the efficiency of CRISPR-induced gene modification in plants, which include the use of optimal transformation and regeneration protocols, the design of appropriate guide RNAs and donor templates and the choice of nucleases and means of delivery. We also present what can be done to orient DNA repair pathways in the target cells, and we show how the moss Physcomitrella patens can be used as a model plant to better understand what DNA repair mechanisms are involved, and how this knowledge could eventually be used to define more performant strategies of CRISPR-induced gene knock-in.

Functional interrogation of non-coding DNA through CRISPR genome editing

Methodologies to interrogate non-coding regions have lagged behind coding regions despite comprising the vast majority of the genome. However, the rapid evolution of clustered regularly interspaced short palindromic repeats (CRISPR)-based genome editing has provided a multitude of novel techniques for laboratory investigation including significant contributions to the toolbox for studying non-coding DNA. CRISPR-mediated loss-of-function strategies rely on direct disruption of the underlying sequence or repression of transcription without modifying the targeted DNA sequence. CRISPR-mediated gain-of-function approaches similarly benefit from methods to alter the targeted sequence through integration of customized sequence into the genome as well as methods to activate transcription. Here we review CRISPR-based loss- and gain-of-function techniques for the interrogation of non-coding DNA.

Marker-free coselection for CRISPR-driven genome editing in human cells

Targeted genome editing enables the creation of bona fide cellular models for biological research and may be applied to human cell-based therapies. Therefore, broadly applicable and versatile methods for increasing its efficacy in cell populations are highly desirable. We designed a simple and robust coselection strategy for enrichment of cells with either nuclease-driven nonhomologous end joining (NHEJ) or homology-directed repair (HDR) events by harnessing the multiplexing capabilities of CRISPR-Cas9 and Cpf1 systems. Selection for dominant alleles of the ubiquitous sodium/potassium pump (Na⁺/K⁺ ATPase) that rendered cells resistant to ouabain was used to enrich for custom genetic modifications at another unlinked locus of interest, thereby effectively increasing the recovery of engineered cells. The process is readily adaptable to transformed and primary cells, including hematopoietic stem and progenitor cells. The use of universal CRISPR reagents and a commercially available small-molecule inhibitor streamlines the incorporation of marker-free genetic changes in human cells.

Precision genome editing in the CRISPR era

With the introduction of precision genome editing using CRISPR-Cas9 technology, we have entered a new era of genetic engineering and gene therapy. With RNA-guided endonucleases, such as Cas9, it is possible to engineer DNA double strand breaks (DSB) at specific genomic loci. DSB repair by the error-prone non-homologous end-joining (NHEJ) pathway can disrupt a target gene by generating insertions and deletions. Alternatively, Cas9-mediated DSBs can be repaired by homology-directed repair (HDR) using an homologous DNA repair template, thus allowing precise gene editing by incorporating genetic changes into the repair template. HDR can introduce gene sequences for protein epitope tags, delete genes, make point mutations, or alter enhancer and promoter activities. In anticipation of adapting this technology for gene therapy in human somatic cells, much focus has been placed on increasing the fidelity of CRISPR-Cas9 and increasing HDR efficiency to improve precision genome editing. In this review, we will discuss applications of CRISPR technology for gene inactivation and genome editing with a focus on approaches to enhancing CRISPR-Cas9-mediated HDR for the generation of cell and animal models, and conclude with a discussion of recent advances and challenges towards the application of this technology for gene therapy in humans.

Functional variomics and network perturbation: connecting genotype to phenotype in cancer

Proteins interact with other macromolecules in complex cellular networks for signal transduction and biological function. In cancer, genetic aberrations have been traditionally thought to disrupt the entire gene function. It has been increasingly appreciated that each mutation of a gene could have a subtle but unique effect on protein function or network rewiring, contributing to diverse phenotypic consequences across cancer patient populations. In this Review, we discuss the current understanding of cancer genetic variants, including the broad spectrum of mutation classes and the wide range of mechanistic effects on gene function in the context of signalling networks. We highlight recent advances in computational and experimental strategies to study the diverse functional and phenotypic consequences of mutations at the base-pair resolution. Such information is crucial to understanding the complex pleiotropic effect of cancer genes and provides a possible link between genotype and phenotype in cancer.

Efficient Precision Genome Editing in iPSCs via Genetic Co-targeting with Selection

Genome editing in induced pluripotent stem cells is currently hampered by the laborious and expensive nature of identifying homology-directed repair (HDR)-modified cells. We present an approach where isolation of cells bearing a selectable, HDR-mediated editing event at one locus enriches for HDR-mediated edits at additional loci. This strategy, called co-targeting with selection, improves the probability of isolating cells bearing HDR-mediated variants and accelerates the production of disease models.

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Increases the efficiency of genome editing in human iPSCs

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Enhances detectability of variants of interest derived by homology-directed repair

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Is a simple, scalable, and adaptable strategy for knocking in variants of interest

Efficient precise knockin with a double cut HDR donor after CRISPR/Cas9-mediated double-stranded DNA cleavage

BACKGROUND

Precise genome editing via homology-directed repair (HDR) after double-stranded DNA (dsDNA) cleavage facilitates functional genomic research and holds promise for gene therapy. However, HDR efficiency remains low in some cell types, including some of great research and clinical interest, such as human induced pluripotent stem cells (iPSCs).

RESULTS

Here, we show that a double cut HDR donor, which is flanked by single guide RNA (sgRNA)-PAM sequences and is released after CRISPR/Cas9 cleavage, increases HDR efficiency by twofold to fivefold relative to circular plasmid donors at one genomic locus in 293 T cells and two distinct genomic loci in iPSCs. We find that a 600 bp homology in both arms leads to high-level genome knockin, with 97-100% of the donor insertion events being mediated by HDR. The combined use of CCND1, a cyclin that functions in G1/S transition, and nocodazole, a G2/M phase synchronizer, doubles HDR efficiency to up to 30% in iPSCs.

CONCLUSIONS

Taken together, these findings provide guidance for the design of HDR donor vectors and the selection of HDR-enhancing factors for applications in genome research and precision medicine.

Organoid technologies meet genome engineering

Three-dimensional (3D) stem cell differentiation cultures recently emerged as a novel model system for investigating human embryonic development and disease progression in vitro, complementing existing animal and two-dimensional (2D) cell culture models. Organoids, the 3D self-organizing structures derived from pluripotent or somatic stem cells, can recapitulate many aspects of structural organization and functionality of their in vivo organ counterparts, thus holding great promise for biomedical research and translational applications. Importantly, faithful recapitulation of disease and development processes relies on the ability to modify the genomic contents in organoid cells. The revolutionary genome engineering technologies, CRISPR/Cas9 in particular, enable investigators to generate various reporter cell lines for prompt validation of specific cell lineages as well as to introduce disease-associated mutations for disease modeling. In this review, we provide historical overviews, and discuss technical considerations, and potential future applications of genome engineering in 3D organoid models.

CRISPR/Cas9-Induced (CTG⋅CAG)n Repeat Instability in the Myotonic Dystrophy Type 1 Locus: Implications for Therapeutic Genome Editing

Myotonic dystrophy type 1 (DM1) is caused by (CTG⋅CAG)n-repeat expansion within the DMPK gene and thought to be mediated by a toxic RNA gain of function. Current attempts to develop therapy for this disease mainly aim at destroying or blocking abnormal properties of mutant DMPK (CUG)n RNA. Here, we explored a DNA-directed strategy and demonstrate that single clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9-cleavage in either its 5' or 3' unique flank promotes uncontrollable deletion of large segments from the expanded trinucleotide repeat, rather than formation of short indels usually seen after double-strand break repair. Complete and precise excision of the repeat tract from normal and large expanded DMPK alleles in myoblasts from unaffected individuals, DM1 patients, and a DM1 mouse model could be achieved at high frequency by dual CRISPR/Cas9-cleavage at either side of the (CTG⋅CAG)n sequence. Importantly, removal of the repeat appeared to have no detrimental effects on the expression of genes in the DM1 locus. Moreover, myogenic capacity, nucleocytoplasmic distribution, and abnormal RNP-binding behavior of transcripts from the edited DMPK gene were normalized. Dual sgRNA-guided excision of the (CTG⋅CAG)n tract by CRISPR/Cas9 technology is applicable for developing isogenic cell lines for research and may provide new therapeutic opportunities for patients with DM1.

Integration and exchange of split dCas9 domains for transcriptional controls in mammalian cells

Programmable and precise regulation of dCas9 functions in response to multiple molecular signals by using synthetic gene circuits will expand the application of the CRISPR-Cas technology. However, the application of CRISPR-Cas therapeutic circuits is still challenging due to the restrictive cargo size of existing viral delivery vehicles. Here, we construct logic AND circuits by integrating multiple split dCas9 domains, which is useful to reduce the size of synthetic circuits. In addition, we engineer sensory switches by exchanging split dCas9 domains, allowing differential regulations on one gene, or activating two different genes in response to cell-type specific microRNAs. Therefore, we provide a valuable split-dCas9 toolkit to engineer complex transcription controls, which may inspire new biomedical applications.

Molecular engineering of Cas9 has the potential to expand the application of CRISPR-Cas technology. Here, Ma et al. show that dCas9 can be split and reconstituted in human cells and use a domain swapping strategy to engineer custom Cas9-based logic circuits and sensory switches.