Control of gene editing by manipulation of DNA repair mechanisms

Topoisomerase Inhibitors and PIM1 Kinase Inhibitors Improve Gene Editing Efficiency Mediated by CRISPR-Cas9 and Homology-Directed Repair

The CRISPR-Cas9 system has emerged as the most prevalent gene editing technology due to its simplicity, high efficiency, and low cost. However, the homology-directed repair (HDR)-mediated gene knock-in in this system suffers from low efficiency, which limits its application in animal model preparation, gene therapy, and agricultural genetic improvement. Here, we report the design and optimization of a simple and efficient reporter-based assay to visualize and quantify HDR efficiency. Through random screening of a small molecule compound library, two groups of compounds, including the topoisomerase inhibitors and PIM1 kinase inhibitors, have been identified to promote HDR. Two representative compounds, etoposide and quercetagetin, also significantly enhance the efficiency of CRISPR-Cas9 and HDR-mediated gene knock-in in mouse embryos. Our study not only provides an assay to screen compounds that may facilitate HDR but also identifies useful tool compounds to facilitate the construction of genetically modified animal models with the CRISPR-Cas9 system.

Allele-specific CRISPR-Cas9 editing of dominant epidermolysis bullosa simplex in human epidermal stem cells

Epidermolysis bullosa simplex (EBS) is a rare skin disease inherited mostly in an autosomal dominant manner. Patients display a skin fragility that leads to blisters and erosions caused by minor mechanical trauma. EBS phenotypic and genotypic variants are caused by genetic defects in intracellular proteins whose function is to provide the attachment of basal keratinocytes to the basement membrane zone and most EBS cases display mutations in keratin 5 (KRT5) and keratin 14 (KRT14) genes. Besides palliative treatments, there is still no long-lasting effective cure to correct the mutant gene and abolish the dominant negative effect of the pathogenic protein over its wild-type counterpart. Here, we propose a molecular strategy for EBS01 patient's keratinocytes carrying a monoallelic c.475/495del21 mutation in KRT14 exon 1. Through the CRISPR-Cas9 system, we perform a specific cleavage only on the mutant allele and restore a normal cellular phenotype and a correct intermediate filament network, without affecting the epidermal stem cell, referred to as holoclones, which play a crucial role in epidermal regeneration.

Genetic manipulation of betta fish

Betta splendens, also known as Siamese fighting fish or "betta," is a freshwater fish species renowned for its astonishing morphological diversity and extreme aggressive behavior. Despite recent advances in our understanding of the genetics and neurobiology of betta, the lack of tools to manipulate their genome has hindered progress at functional and mechanistic levels. In this study, we outline the use of three genetic manipulation technologies, which we have optimized for use in betta: CRISPR/Cas9-mediated knockout, CRISPR/Cas9-mediated knockin, and Tol2-mediated transgenesis. We knocked out three genes: alkal2l, bco1l, and mitfa, and analyzed their effects on viability and pigmentation. Furthermore, we knocked in a fluorescent protein into the mitfa locus, a proof-of-principle experiment of this powerful technology in betta. Finally, we used Tol2-mediated transgenesis to create fish with ubiquitous expression of GFP, and then developed a bicistronic plasmid with heart-specific expression of a red fluorescent protein to serve as a visible marker of successful transgenesis. Our work highlights the potential for the genetic manipulation of betta, providing valuable resources for the effective use of genetic tools in this animal model.

Genetic manipulation of betta fish

Betta splendens , also known as Siamese fighting fish or 'betta', are renowned for their astonishing morphological diversity and extreme aggressive behavior. Despite recent advances in our understanding of the genetics and neurobiology of betta, the lack of tools to manipulate their genome has hindered progress at functional and mechanistic levels. In this study, we outline the use of three genetic manipulation technologies, which we have optimized for use in betta: CRISPR/Cas9-mediated knockout, CRISPR/Cas9-mediated knockin, and Tol2-mediated transgenesis. We knocked out three genes: alkal2l, bco1l , and mitfa , and analyzed their effects on viability and pigmentation. Furthermore, we successfully knocked in a fluorescent protein into the mitfa locus, a proof-of-principle experiment of this powerful technology in betta. Finally, we used Tol2-mediated transgenesis to create fish with ubiquitous expression of GFP, and then developed a bicistronic plasmid with heart-specific expression of a red fluorescent protein to serve as a visible marker of successful transgenesis. Our work highlights the potential for the genetic manipulation of betta, providing valuable resources for the effective use of genetic tools in this animal model.

Genome Editing Using Cas9 Ribonucleoprotein Is Effective for Introducing PDGFRA Variant in Cultured Human Glioblastoma Cell Lines

Many variants of uncertain significance (VUS) have been detected in clinical cancer cases using next-generation sequencing-based cancer gene panel analysis. One strategy for the elucidation of VUS is the functional analysis of cultured cancer cell lines that harbor targeted gene variants using genome editing. Genome editing is a powerful tool for creating desired gene alterations in cultured cancer cell lines. However, the efficiency of genome editing varies substantially among cell lines of interest. We performed comparative studies to determine the optimal editing conditions for the introduction of platelet-derived growth factor receptor alpha (PDGFRA) variants in human glioblastoma multiforme (GBM) cell lines. After monitoring the copy numbers of PDGFRA and the expression level of the PDGFRα protein, four GBM cell lines (U-251 MG, KNS-42, SF126, and YKG-1 cells) were selected for the study. To compare the editing efficiency in these GBM cell lines, the modes of clustered regularly interspaced short palindromic repeat (CRISPR)-associated protein 9 (Cas9) delivery (plasmid vs. ribonucleoprotein (RNP)), methods of transfection (lipofection vs. electroporation), and usefulness of cell sorting were then evaluated. Herein, we demonstrated that electroporation-mediated transfer of Cas9 with single-guide RNA (Cas9 RNP complex) could sufficiently edit a target nucleotide substitution, irrespective of cell sorting. As the Cas9 RNP complex method showed a higher editing efficiency than the Cas9 plasmid lipofection method, it was the optimal method for single-nucleotide editing in human GBM cell lines under our experimental conditions.

Multiplex CRISPR/Cas9-mediated genome editing to address drought tolerance in wheat

Genome editing tools have rapidly been adopted by plant scientists for crop improvement. Genome editing using a multiplex sgRNA-CRISPR/Cas9 genome editing system is a useful technique for crop improvement in monocot species. In this study, we utilized precise gene editing techniques to generate wheat 3'(2'), 5'-bisphosphate nucleotidase (TaSal1) mutants using a multiplex sgRNA-CRISPR/Cas9 genome editing system. Five active TaSal1 homologous genes were found in the genome of Giza168 in addition to another apparently inactive gene on chromosome 4A. Three gRNAs were designed and used to target exons 4, 5 and 7 of the five wheat TaSal1 genes. Among the 120 Giza168 transgenic plants, 41 lines exhibited mutations and produced heritable TaSal1 mutations in the M1 progeny and 5 lines were full 5 gene knock-outs. These mutant plants exhibit a rolled-leaf phenotype in young leaves and bended stems, but there were no significant changes in the internode length and width, leaf morphology, and stem shape. Anatomical and scanning electron microscope studies of the young leaves of mutated TaSal1 lines showed closed stomata, increased stomata width and increase in the size of the bulliform cells. Sal1 mutant seedlings germinated and grew better on media containing polyethylene glycol than wildtype seedlings. Our results indicate that the application of the multiplex sgRNA-CRISPR/Cas9 genome editing is efficient tool for mutating more multiple TaSal1 loci in hexaploid wheat.

Quantitative analysis of CRISPR/Cas9-mediated provirus deletion in blue egg layer chicken PGCs by digital PCR

Primordial germ cells (PGCs), the precursors of sperm and oocytes, pass on the genetic material to the next generation. The previously established culture system of chicken PGCs holds many possibilities for functional genomics studies and the rapid introduction of desired traits. Here, we established a CRISPR/Cas9-mediated genome editing protocol for the genetic modification of PGCs derived from chickens with blue eggshell color. The sequence targeted in the present report is a provirus (EAV-HP) insertion in the 5'-flanking region of the SLCO1B3 gene on chromosome 1 in Araucana chickens, which is supposedly responsible for the blue eggshell color. We designed pairs of guide RNAs (gRNAs) targeting the entire 4.2 kb provirus region. Following transfection of PGCs with the gRNA, genomic DNA was isolated and analyzed by mismatch cleavage assay (T7EI). For absolute quantification of the targeting efficiencies in homozygous blue-allele bearing PGCs a digital PCR was established, which revealed deletion efficiencies of 29% when the wildtype Cas9 was used, and 69% when a high-fidelity Cas9 variant was employed. Subsequent single cell dilutions of edited PGCs yielded 14 cell clones with homozygous deletion of the provirus. A digital PCR assay proved the complete absence of this provirus in cell clones. Thus, we demonstrated the high efficiency of the CRISPR/Cas9 system in introducing a large provirus deletion in chicken PGCs. Our presented workflow is a cost-effective and rapid solution for screening the editing success in transfected PGCs.

Integrase deficient lentiviral vector: prospects for safe clinical applications

HIV-1 derived lentiviral vector is an efficient transporter for delivering desired genetic materials into the targeted cells among many viral vectors. Genetic material transduced by lentiviral vector is integrated into the cell genome to introduce new functions, repair defective cell metabolism, and stimulate certain cell functions. Various measures have been administered in different generations of lentiviral vector systems to reduce the vector's replicating capabilities. Despite numerous demonstrations of an excellent safety profile of integrative lentiviral vectors, the precautionary approach has prompted the development of integrase-deficient versions of these vectors. The generation of integrase-deficient lentiviral vectors by abrogating integrase activity in lentiviral vector systems reduces the rate of transgenes integration into host genomes. With this feature, the integrase-deficient lentiviral vector is advantageous for therapeutic implementation and widens its clinical applications. This short review delineates the biology of HIV-1-erived lentiviral vector, generation of integrase-deficient lentiviral vector, recent studies involving integrase-deficient lentiviral vectors, limitations, and prospects for neoteric clinical use.

Transcription-coupled donor DNA expression increases homologous recombination for efficient genome editing

Genomes can be edited by homologous recombination stimulated by CRISPR/Cas9 [clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated peptide 9]-induced DNA double-strand breaks. However, this approach is inefficient for inserting or deleting long fragments in mammalian cells. Here, we describe a simple genome-editing method, termed transcription-coupled Cas9-mediated editing (TEd), that can achieve higher efficiencies than canonical Cas9-mediated editing (CEd) in deleting genomic fragments, inserting/replacing large DNA fragments and introducing point mutations into mammalian cell lines. We also found that the transcription on DNA templates is crucial for the promotion of homology-directed repair, and that tethering transcripts from TEd donors to targeted sites further improves editing efficiency. The superior efficiency of TEd for the insertion and deletion of long DNA fragments expands the applications of CRISPR for editing mammalian genomes.

New developments in the molecular treatment of ichthyosis: review of the literature

Ichthyosis covers a wide spectrum of diseases affecting the cornification of the skin. In recent years, new advances in understanding the pathophysiology of ichthyosis have been made. This knowledge, combined with constant development of pathogenesis-based therapies, such as protein replacement therapy and gene therapy, are rather promising for patients with inherited skin diseases. Several ongoing trials are investigating the potency of these new approaches and various studies have already been published. Furthermore, a lot of case series report that biological therapeutics are effective treatment options, mainly for Netherton syndrome and autosomal recessive congenital ichthyosis. It is expected that some of these new therapies will prove their efficacy and will be incorporated in the treatment of ichthyosis.

From DNA break repair pathways to CRISPR/Cas-mediated gene knock-in methods

Various DNA breaks created via programmable CRISPR/Cas9 nuclease activity results in different intracellular DNA break repair pathways. Based on the cellular repair pathways, CRISPR-based gene knock-in methods can be categorized into two major strategies: 1) Homology-independent strategies which are targeted insertion events based on non-homologous end joining, and 2) Homology-dependent strategies which are targeted insertion events based on the homology-directed repair. This review elaborates on various gene knock-in methods in mammalian cells using the CRISPR/Cas9 system and in sync with DNA-break repair pathways. Gene knock-in methods are applied in functional genomics and gene therapy. To compensate or correct genetic defects, different CRISPR-based gene knock-in strategies can be used. Thus, researchers need to make a conscious decision about the most suitable knock-in method. For a successful gene-targeted insertion, some determinant factors should be considered like cell cycle, dominant DNA repair pathway, size of insertions, and donor properties. In this review, different aspects of each gene knock-in strategy are discussed to provide a framework for choosing the most appropriate gene knock-in method in different applications.

Endogenous protein tagging in medaka using a simplified CRISPR/Cas9 knock-in approach

The CRISPR/Cas9 system has been used to generate fluorescently labelled fusion proteins by homology-directed repair in a variety of species. Despite its revolutionary success, there remains an urgent need for increased simplicity and efficiency of genome editing in research organisms. Here, we establish a simplified, highly efficient, and precise strategy for CRISPR/Cas9-mediated endogenous protein tagging in medaka (Oryzias latipes). We use a cloning-free approach that relies on PCR-amplified donor fragments containing the fluorescent reporter sequences flanked by short homology arms (30–40 bp), a synthetic single-guide RNA and Cas9 mRNA. We generate eight novel knock-in lines with high efficiency of F0 targeting and germline transmission. Whole genome sequencing results reveal single-copy integration events only at the targeted loci. We provide an initial characterization of these fusion protein lines, significantly expanding the repertoire of genetic tools available in medaka. In particular, we show that the mScarlet-pcna line has the potential to serve as an organismal-wide label for proliferative zones and an endogenous cell cycle reporter.

A non-viral and selection-free COL7A1 HDR approach with improved safety profile for dystrophic epidermolysis bullosa

Gene editing via homology-directed repair (HDR) currently comprises the best strategy to obtain perfect corrections for pathogenic mutations of monogenic diseases, such as the severe recessive dystrophic form of the blistering skin disease epidermolysis bullosa (RDEB). Limitations of this strategy, in particular low efficiencies and off-target effects, hinder progress toward clinical applications. However, the severity of RDEB necessitates the development of efficient and safe gene-editing therapies based on perfect repair. To this end, we sought to assess the corrective efficiencies following optimal Cas9 nuclease and nickase-based COL7A1-targeting strategies in combination with single- or double-stranded donor templates for HDR at the COL7A1 mutation site. We achieved HDR-mediated correction efficiencies of up to 21% and 10% in primary RDEB keratinocytes and fibroblasts, respectively, as analyzed by next-generation sequencing, leading to full-length type VII collagen restoration and accurate deposition within engineered three-dimensional (3D) skin equivalents (SEs). Extensive on- and off-target analyses confirmed that the combined treatment of paired nicking and single-stranded oligonucleotides constituted a highly efficient COL7A1-editing strategy, associated with a significantly improved safety profile. Our findings, therefore, represent a further advancement in the field of traceless genome editing for genodermatoses.

Base Editing in Plants: Applications, Challenges, and Future Prospects

The ability to create targeted modifications in the genomes of plants using genome editing technologies has revolutionized research in crop improvement in the current dispensation of molecular biology. This technology has attracted global attention and has been employed in functional analysis studies in crop plants. Since many important agronomic traits are confirmed to be determined by single-nucleotide polymorphisms, improved crop varieties could be developed by the programmed and precise conversion of targeted single bases in the genomes of plants. One novel genome editing approach which serves for this purpose is base editing. Base editing directly makes targeted and irreversible base conversion without creating double-strand breaks (DSBs). This technology has recently gained quick acceptance and adaptation because of its precision, simplicity, and multiplex capabilities. This review focuses on generating different base-editing technologies and how efficient they are in editing nucleic acids. Emphasis is placed on the exploration and applications of these base-editing technologies to enhance crop production. The review also highlights the drawbacks and the prospects of this new technology.

Chlamydomonas POLQ is necessary for CRISPR/Cas9-mediated gene targeting

The use of CRISPR/Cas endonucleases has revolutionized gene editing techniques for research on Chlamydomonas reinhardtii. To better utilize the CRISPR/Cas system, it is essential to develop a more comprehensive understanding of the DNA repair pathways involved in genome editing. In this study, we have analyzed contributions from canonical KU80/KU70-dependent nonhomologous end-joining (cNHEJ) and DNA polymerase theta (POLQ)-mediated end joining on SpCas9-mediated untemplated mutagenesis and homology-directed repair (HDR)/gene inactivation in Chlamydomonas. Using CRISPR/SpCas9 technology, we generated DNA repair-defective mutants ku80, ku70, polQ for gene targeting experiments. Our results show that untemplated repair of SpCas9-induced double strand breaks results in mutation spectra consistent with an involvement of both KU80/KU70 and POLQ. In addition, the inactivation of POLQ was found to negatively affect HDR of the inactivated paromomycin-resistant mut-aphVIII gene when donor single-stranded oligos were used. Nevertheless, mut-aphVIII was still repaired by homologous recombination in these mutants. POLQ inactivation suppressed random integration of transgenes co-transformed with the donor ssDNA. KU80 deficiency did not affect these events but instead was surprisingly found to stimulate HDR/gene inactivation. Our data suggest that in Chlamydomonas, POLQ is the main contributor to CRISPR/Cas-induced HDR and random integration of transgenes, whereas KU80/KU70 potentially plays a secondary role. We expect our results will lead to improvement of genome editing in C. reinhardtii and can be used for future development of algal biotechnology.

Present and future prospects for wheat improvement through genome editing and advanced technologies

Wheat (Triticum aestivum, 2n = 6x = 42, AABBDD) is one of the most important staple food crops in the world. Despite the fact that wheat production has significantly increased over the past decades, future wheat production will face unprecedented challenges from global climate change, increasing world population, and water shortages in arid and semi-arid lands. Furthermore, excessive applications of diverse fertilizers and pesticides are exacerbating environmental pollution and ecological deterioration. To ensure global food and ecosystem security, it is essential to enhance the resilience of wheat production while minimizing environmental pollution through the use of cutting-edge technologies. However, the hexaploid genome and gene redundancy complicate advances in genetic research and precision gene modifications for wheat improvement, thus impeding the breeding of elite wheat cultivars. In this review, we first introduce state-of-the-art genome-editing technologies in crop plants, especially wheat, for both functional genomics and genetic improvement. We then outline applications of other technologies, such as GWAS, high-throughput genotyping and phenotyping, speed breeding, and synthetic biology, in wheat. Finally, we discuss existing challenges in wheat genome editing and future prospects for precision gene modifications using advanced genome-editing technologies. We conclude that the combination of genome editing and other molecular breeding strategies will greatly facilitate genetic improvement of wheat for sustainable global production.

Targeted integration into pseudo attP sites of CHO cells using CRISPR/Cas9

Chinese hamster ovary (CHO) cells are regarded as a prominent host for manufacturing therapeutic proteins. Although conventional strategies for generating recombinant proteins in CHO cells depend on the random integration of a gene of interest (GOI), these established techniques occasionally result in genetically heterogeneous cell lines, which causes diminished expression of the recombinant proteins in the long run. Production instability can be reduced by SSI and creates stable cell lines with a consistent expression of the GOI. In this experiment, we demonstrate the targeted incorporation of a reporter cassette in two PhiC31 pseudo attP sites of CHO cells exploiting the homology-directed repair (HDR) generated by the CRISPR/Cas9 platform. Genes encoding GFP and puromycin resistance marker were precisely inserted into these loci via CRISPR/Cas9. Stable cell lines were suitably produced following antibiotic selection. Junction PCR and fluorescence assay determined targeted integration and expression homogeneity of the reporter cassette, respectively. Taken together, our results indicate the possibility of these two PhiC31 pseudo attP sites as the target sites for site-specific integration of a transgene mediated by CRISPR/Cas9. Furthermore, higher knock-in efficiency and expression homogeneity was observed in the pseudo attP site associated with chromosome 6 compared to the pseudo attP site from chromosome 3.

An Optimized Preparation Method for Long ssDNA Donors to Facilitate Quick Knock-In Mouse Generation

Fluorescent reporter mouse lines and Cre/Flp recombinase driver lines play essential roles in investigating various molecular functions in vivo. Now that applications of the CRISPR/Cas9 genome-editing system to mouse fertilized eggs have drastically accelerated these knock-in mouse generations, the next need is to establish easier, quicker, and cheaper methods for knock-in donor preparation. Here, we reverify and optimize the phospho-PCR method to obtain highly pure long single-stranded DNAs (ssDNAs) suitable for knock-in mouse generation via genome editing. The sophisticated sequential use of two exonucleases, in which double-stranded DNAs (dsDNAs) amplified by a pair of 5′-phosphorylated primer and normal primer are digested by Lambda exonuclease to yield ssDNA and the following Exonuclease III treatment degrades the remaining dsDNAs, enables much easier long ssDNA productions without laborious gel extraction steps. By microinjecting these donor DNAs along with CRISPR/Cas9 components into mouse zygotes, we have effectively generated fluorescent reporter lines and recombinase drivers. To further broaden the applicability, we have prepared long ssDNA donors in higher concentrations and electroporated them into mouse eggs to successfully obtain knock-in embryos. This classical yet improved method, which is regaining attention on the progress of CRISPR/Cas9 development, shall be the first choice for long donor DNA preparation, and the resulting knock-in lines could accelerate life science research.

Stable Cells with NF-κB-ZsGreen Fused Genes Created by TALEN Editing and Homology Directed Repair for Screening Anti-inflammation Drugs

Background

NF-κB is a sequence-specific DNA-binding transcription factor that plays key roles in inflammation and cancer. It is well known that NF-κB is over-activated in these diseases. NF-κB inhibitors are therefore developed as promising drugs for these diseases. However, finding NF-κB inhibitors is dependent on effective screening platforms.

Methods

For providing an easy and visualizable tool for screening NF-κB inhibitors, and other NF-κB-related studies, this study edited all five genes of NF-κB family (RELA, RELB, CREL, NF-κB1, NF-κB2) in three different cell lines (293T, HepG2, and PANC1) with both TALEN and CRISPR. The edited NF-κB genes were repaired by homology-dependent repair using a linear homologous donor containing ZsGreen coding sequence. The edit efficiency was thus directly evaluated by detecting cellular fluorescence. The editing efficiency was also confirmed by PCR detection of NF-κB-ZsGreen fused genes.

Results

It was found that all genes were more efficiently edited by TALEN in all cells than CRISPR. The positive cells were then isolated from the TALEN-edited cell pool by flow cytometry. The purified positive cells were finally evaluated by regulating NF-κB activity with a known NF-κB inhibitor, BAY 11-7082, and an NF-κB-targeting artificial microRNA, miR533. The results revealed that all the labeled NF-κB genes responded well to the two kinds of NF-κB activity regulators in all cell lines.

Conclusion

This study thus obtained 15 cell lines with NF-κB-ZsGreen fused genes, which provide an easy and visualizable tool for screening NF-κB inhibitors and other NF-κB-related studies.

A CRISPR/Cas9-Mediated, Homology-Independent Tool Developed for Targeted Genome Integration in Yarrowia lipolytica

Yarrowia lipolytica has been extensively used to produce essential chemicals and enzymes. As in most other eukaryotes, nonhomologous end joining (NHEJ) is the major repair pathway for DNA double-strand breaks in Y. lipolytica Although numerous studies have attempted to achieve targeted genome integration through homologous recombination (HR), this process requires the construction of homologous arms, which is time-consuming. This study aimed to develop a homology-independent and CRISPR/Cas9-mediated targeted genome integration tool in Y. lipolytica Through optimization of the cleavage efficiency of Cas9, targeted integration of a hyg fragment was achieved with 12.9% efficiency, which was further improved by manipulation of the fidelity of NHEJ repair, the cell cycle, and the integration sites. Thus, the targeted integration rate reached 55% through G₁ phase synchronization. This tool was successfully applied for the rapid verification of intronic promoters and iterative integration of four genes in the pathway for canthaxanthin biosynthesis. This homology-independent integration tool does not require homologous templates and selection markers and achieves one-step targeted genome integration of the 8,417-bp DNA fragment, potentially replacing current HR-dependent genome-editing methods for Y. lipolytica IMPORTANCE This study describes the development and optimization of a homology-independent targeted genome integration tool mediated by CRISPR/Cas9 in Yarrowia lipolytica This tool does not require the construction of homologous templates and can be used to rapidly verify genetic elements and to iteratively integrate multiple-gene pathways in Y. lipolytica This tool may serve as a potential supplement to current HR-dependent genome-editing methods for eukaryotes.

One-step generation of a targeted knock-in calf using the CRISPR-Cas9 system in bovine zygotes

Background

The homologous recombination (HR) pathway is largely inactive in early embryos prior to the first cell division, making it difficult to achieve targeted gene knock-ins. The homology-mediated end joining (HMEJ)-based strategy has been shown to increase knock-in efficiency relative to HR, non-homologous end joining (NHEJ), and microhomology-mediated end joining (MMEJ) strategies in non-dividing cells.

Results

By introducing gRNA/Cas9 ribonucleoprotein complex and a HMEJ-based donor template with 1 kb homology arms flanked by the H11 safe harbor locus gRNA target site, knock-in rates of 40% of a 5.1 kb bovine sex-determining region Y (SRY)-green fluorescent protein (GFP) template were achieved in Bos taurus zygotes. Embryos that developed to the blastocyst stage were screened for GFP, and nine were transferred to recipient cows resulting in a live phenotypically normal bull calf. Genomic analyses revealed no wildtype sequence at the H11 target site, but rather a 26 bp insertion allele, and a complex 38 kb knock-in allele with seven copies of the SRY-GFP template and a single copy of the donor plasmid backbone. An additional minor 18 kb allele was detected that looks to be a derivative of the 38 kb allele resulting from the deletion of an inverted repeat of four copies of the SRY-GFP template.

Conclusion

The allelic heterogeneity in this biallelic knock-in calf appears to have resulted from a combination of homology directed repair, homology independent targeted insertion by blunt-end ligation, NHEJ, and rearrangement following editing of the gRNA target site in the donor template.

This study illustrates the potential to produce targeted gene knock-in animals by direct cytoplasmic injection of bovine embryos with gRNA/Cas9, although further optimization is required to ensure a precise single-copy gene integration event.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12864-021-07418-3.

Advances in gene editing strategies for epidermolysis bullosa

Epidermolysis bullosa represents a monogenetic disease comprising a variety of heterogeneous mutations in at least 16 genes encoding structural proteins crucial for skin integrity. Due to well-defined mutations but still lacking causal treatment options for the disease, epidermolysis bullosa represents an ideal candidate for gene therapeutic interventions. Recent developments and improvements in the genome editing field have paved the way for the translation of various gene repair strategies into the clinic. With the ability to accurately predict and monitor targeting events within the human genome, the translation might soon be possible. Here, we describe current advancements in the genome editing field for epidermolysis bullosa, along with a discussion of aspects and strategies for precise and personalized gene editing-based medicine, in order to develop efficient and safe ex vivo as well as in vivo genome editing therapies for epidermolysis bullosa patients in the future.

DGK and DZHK position paper on genome editing: basic science applications and future perspective

For a long time, gene editing had been a scientific concept, which was limited to a few applications. With recent developments, following the discovery of TALEN zinc-finger endonucleases and in particular the CRISPR/Cas system, gene editing has become a technique applicable in most laboratories. The current gain- and loss-of function models in basic science are revolutionary as they allow unbiased screens of unprecedented depth and complexity and rapid development of transgenic animals. Modifications of CRISPR/Cas have been developed to precisely interrogate epigenetic regulation or to visualize DNA complexes. Moreover, gene editing as a clinical treatment option is rapidly developing with first trials on the way. This article reviews the most recent progress in the field, covering expert opinions gathered during joint conferences on genome editing of the German Cardiac Society (DGK) and the German Center for Cardiovascular Research (DZHK). Particularly focusing on the translational aspect and the combination of cellular and animal applications, the authors aim to provide direction for the development of the field and the most frequent applications with their problems.

A Universal System of CRISPR/Cas9-Mediated Gene Targeting Using All-in-One Vector in Plants

Homologous recombination-mediated genome editing, also called gene targeting (GT), is an essential technique that allows precise modification of a target sequence, including introduction of point mutations, knock-in of a reporter gene, and/or swapping of a functional domain. However, due to its low frequency, it has been difficult to establish GT approaches that can be applied widely to a large number of plant species. We have developed a simple and universal clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9)-mediated DNA double-strand break (DSB)-induced GT system using an all-in-one vector comprising a CRISPR/Cas9 expression construct, selectable marker, and GT donor template. This system enabled introduction of targeted point mutations with non-selectable traits into several target genes in both rice and tobacco. Since it was possible to evaluate the GT frequency on endogenous target genes precisely using this system, we investigated the effect of treatment with Rad51-stimulatory compound 1 (RS-1) on the frequency of DSB-induced GT. GT frequency was slightly, but consistently, improved by RS-1 treatment in both target plants.

Harnessing endogenous repair mechanisms for targeted gene knock-in of bovine embryos

Introducing useful traits into livestock breeding programs through gene knock-ins has proven challenging. Typically, targeted insertions have been performed in cell lines, followed by somatic cell nuclear transfer cloning, which can be inefficient. An alternative is to introduce genome editing reagents and a homologous recombination (HR) donor template into embryos to trigger homology directed repair (HDR). However, the HR pathway is primarily restricted to actively dividing cells (S/G2-phase) and its efficiency for the introduction of large DNA sequences in zygotes is low. The homology-mediated end joining (HMEJ) approach has been shown to improve knock-in efficiency in non-dividing cells and to harness HDR after direct injection of embryos. The knock-in efficiency for a 1.8 kb gene was contrasted when combining microinjection of a gRNA/Cas9 ribonucleoprotein complex with a traditional HR donor template or an HMEJ template in bovine zygotes. The HMEJ template resulted in a significantly higher rate of gene knock-in as compared to the HR template (37.0% and 13.8%; P < 0.05). Additionally, more than a third of the knock-in embryos (36.9%) were non-mosaic. This approach will facilitate the one-step introduction of gene constructs at a specific location of the bovine genome and contribute to the next generation of elite cattle.

Adeno-associated virus-mediated gene delivery promotes S-phase entry-independent precise targeted integration in cardiomyocytes

Post-mitotic cardiomyocytes have been considered to be non-permissive to precise targeted integration including homology-directed repair (HDR) after CRISPR/Cas9 genome editing. Here, we demonstrate that direct delivery of large amounts of transgene encoding guide RNA (gRNA) and repair template DNA via intra-ventricular injection of adeno-associated virus (AAV) promotes precise targeted genome replacement in adult murine cardiomyocytes expressing Cas9. Neither systemic injection of AAV nor direct injection of adenovirus promotes targeted integration, suggesting that high copy numbers of single-stranded transgenes are required in cardiomyocytes. Notably, AAV-mediated targeted integration in cardiomyocytes both in vitro and in vivo depends on the Fanconi anemia pathway, a key component of the single-strand template repair mechanism. In human cardiomyocytes differentiated from induced pluripotent stem cells, AAV-mediated targeted integration fluorescently labeled Mlc2v protein after differentiation, independently of DNA synthesis, and enabled real-time detection of sarcomere contraction in monolayered beating cardiomyocytes. Our findings provide a wide range of applications for targeted genome replacement in non-dividing cardiomyocytes.

Rapid Tagging of Human Proteins with Fluorescent Reporters by Genome Engineering using Double-Stranded DNA Donors

Tagging proteins with fluorescent reporters such as green fluorescent protein (GFP) is a powerful method to determine protein localization, especially when proteins are tagged in the endogenous context to preserve native genomic regulation. However, insertion of fluorescent reporters into the genomes of mammalian cells has required the construction of plasmids containing selection markers and/or extended sequences homologous to the site of insertion (homology arms). Here we describe a streamlined protocol that eliminates all cloning steps by taking advantage of the high propensity of linear DNAs to engage in homology‐directed repair of DNA breaks induced by the Cas9 RNA‐guided endonuclease. The protocol uses PCR amplicons, or synthetic gene fragments, with short homology arms (30‐40 bp) to insert fluorescent reporters at specific genomic locations. The linear DNAs are introduced into cells with preassembled Cas9‐crRNA‐tracrRNA complexes using one of two transfection procedures, nucleofection or lipofection. The protocol can be completed under a week, with efficiencies ranging from 0.5% to 20% of transfected cells depending on the locus targeted. © 2019 The Authors.

Variability in Genome Editing Outcomes: Challenges for Research Reproducibility and Clinical Safety

Genome editing tools have already revolutionized biomedical research and are also expected to have an important impact in the clinic. However, their extensive use in research has revealed much unpredictability, both off and on target, in the outcome of their application. We discuss the challenges associated with this unpredictability, both for research and in the clinic. For the former, an extensive validation of the model is essential. For the latter, potential unpredicted activity does not preclude the use of these tools but requires that molecular evidence to underpin the relevant risk:benefit evaluation is available. Safe and successful clinical application will also depend on the mode of delivery and the cellular context.

Gene technologies in weed management: a technical feasibility analysis

With the advent of new genetic technologies such as gene silencing and gene drive, efforts to develop additional management tools for weed management is gaining significant momentum. These technologies promise novel ways to develop sustainable weed control options because gene silencing can switch-off genes mediating adaptation (e.g. growth, herbicide resistance), and gene drive can be used to spread modified traits and to engineer wild populations with reduced fitness. However, applying gene silencing and/or gene drive is expected to be inherently complex as their application is constrained by several methodological and technological difficulties. In this review we explore the challenges of these technologies, and discuss strategies and resources accessible to accelerate the development of gene-tech based tools for weed management. We also highlight how gene technologies can be integrated into existing management tactics such as classical biological control, and their possible interactions.

Context-Dependent Strategies for Enhanced Genome Editing of Genodermatoses

The skin provides direct protection to the human body from assault by the harsh external environment. The crucial function of this organ is significantly disrupted in genodermatoses patients. Genodermatoses comprise a heterogeneous group of largely monogenetic skin disorders, typically involving mutations in genes encoding structural proteins. Therapeutic options for this debilitating group of diseases, including epidermolysis bullosa, primarily consist of wound management. Genome editing approaches co-opt double-strand break repair pathways to introduce desired sequence alterations at specific loci. Rapid advances in genome editing technologies have the potential to propel novel genetic therapies into the clinic. However, the associated phenotypes of many mutations may be treated via several genome editing strategies. Therefore, for potential clinical applications, implementation of efficient approaches based upon mutation, gene and disease context is necessary. Here, we describe current genome editing approaches for the treatment of genodermatoses, along with a discussion of the optimal strategy for each genetic context, in order to achieve enhanced genome editing approaches.

Precise gene replacement in plants through CRISPR/Cas genome editing technology: current status and future perspectives

CRISPR/Cas, as a simple, versatile, robust and cost-effective system for genome manipulation, has dominated the genome editing field over the past few years. The application of CRISPR/Cas in crop improvement is particularly important in the context of global climate change, as well as diverse agricultural, environmental and ecological challenges. Various CRISPR/Cas toolboxes have been developed and allow for targeted mutagenesis at specific genome loci, transcriptome regulation and epigenome editing, base editing, and precise targeted gene/allele replacement or tagging in plants. In particular, precise replacement of an existing allele with an elite allele in a commercial variety through homology-directed repair (HDR) is a holy grail in genome editing for crop improvement as it has been very difficult, laborious and time-consuming to introgress the elite alleles into commercial varieties without any linkage drag from parental lines within a few generations in crop breeding practice. However, it still remains very challenging in crop plants. This review intends to provide an informative summary of the latest development and breakthroughs in gene replacement using CRISPR/Cas technology, with a focus on achievements, potential mechanisms and future perspectives in plant biological science as well as crop improvement.

The Problem of the Low Rates of CRISPR/Cas9-Mediated Knock-ins in Plants: Approaches and Solutions

The main number of genome editing events in plant objects obtained during the last decade with the help of specific nucleases zinc finger (ZFN), transcription activator-like effector nucleases (TALEN), and clustered regularly interspaced short palindromic repeats (CRISPR)/Cas are the microindels causing frameshift and subsequent gene knock-out. The knock-ins of genes or their parts, i.e., the insertion of them into a target genome region, are between one and two orders of magnitude less frequent. First and foremost, this is associated with the specific features of the repair systems of higher eukaryotes and the availability of the donor template in accessible proximity during double-strand break (DSB) repair. This review briefs the main repair pathways in plants according to the aspect of their involvement in genome editing. The main methods for increasing the frequency of knock-ins are summarized both along the homologous recombination pathway and non-homologous end joining, which can be used for plant objects.

Congenital eye anomalies: More mosaic than thought?

The eye is a sensory organ that primarily captures light and provides the sense of sight, as well as delivering non-visual light information involving biological rhythms and neurophysiological activities to the brain. Since the early 1990s, rapid advances in molecular biology have enabled the identification of developmental genes, genes responsible for human congenital diseases, and relevant genes of mutant animals with various anomalies. In this review, we first look at the development of the eye, and we highlight seminal reports regarding archetypal gene defects underlying three developmental ocular disorders in humans: (1) holoprosencephaly (HPE), with cyclopia being exhibited in the most severe cases; (2) microphthalmia, anophthalmia, and coloboma (MAC) phenotypes; and (3) anterior segment dysgenesis (ASDG), known as Peters anomaly and its related disorders. The recently developed methods, such as next-generation sequencing and genome editing techniques, have aided the discovery of gene mutations in congenital eye diseases and gene functions in normal eye development. Finally, we discuss Pax6-genome edited mosaic eyes and propose that somatic mosaicism in developmental gene mutations should be considered a causal factor for variable phenotypes, sporadic cases, and de novo mutations in human developmental disorders.

Making ends meet: targeted integration of DNA fragments by genome editing

Targeted insertion of large pieces of DNA is an important goal of genetic engineering. However, this goal has been elusive since classical methods for homology-directed repair are inefficient and often not feasible in many systems. Recent advances are described here that enable site-specific genomic insertion of relatively large DNA with much improved efficiency. Using the preferred repair pathway in the cell of nonhomologous end-joining, DNA of up to several kb could be introduced with remarkably good precision by the methods of HITI and ObLiGaRe with an efficiency up to 30-40%. Recent advances utilizing homology-directed repair (methods of PITCh; short homology arms including ssODN; 2H2OP) have significantly increased the efficiency for DNA insertion, often to 40-50% or even more depending on the method and length of DNA. The remaining challenges of integration precision and off-target site insertions are summarized. Overall, current advances provide major steps forward for site-specific insertion of large DNA into genomes from a broad range of cells and organisms.

Precision gene editing technology and applications in nephrology

The expanding field of precision gene editing is empowering researchers to directly modify DNA. Gene editing is made possible using synonymous technologies: a DNA-binding platform to molecularly locate user-selected genomic sequences and an associated biochemical activity that serves as a functional editor. The advent of accessible DNA-targeting molecular systems, such as zinc-finger nucleases, transcription activator-like effectors (TALEs) and CRISPR-Cas9 gene editing systems, has unlocked the ability to target nearly any DNA sequence with nucleotide-level precision. Progress has also been made in harnessing endogenous DNA repair machineries, such as non-homologous end joining, homology-directed repair and microhomology-mediated end joining, to functionally manipulate genetic sequences. As understanding of how DNA damage results in deletions, insertions and modifications increases, the genome becomes more predictably mutable. DNA-binding platforms such as TALEs and CRISPR can also be used to make locus-specific epigenetic changes and to transcriptionally enhance or suppress genes. Although many challenges remain, the application of precision gene editing technology in the field of nephrology has enabled the generation of new animal models of disease as well as advances in the development of novel therapeutic approaches such as gene therapy and xenotransplantation.

Formation and nucleolytic processing of Cas9-induced DNA breaks in human cells quantified by droplet digital PCR

Cas9 endonuclease from S. pyogenes is widely used to induce controlled double strand breaks (DSB) at desired genomic loci for gene editing. Here, we describe a droplet digital PCR (ddPCR) method to precisely quantify the kinetic of formation and 5'-end nucleolytic processing of Cas9-induced DSB in different human cells lines. Notably, DSB processing is a finely regulated process, which dictates the choice between non-homologous end joining (NHEJ) and homology directed repair (HDR). This step of DSB repair is also a relevant point to be taken into consideration to improve Cas9-mediated technology. Indeed, by this protocol, we show that processing of Cas9-induced DSB is impaired by CTIP or BRCA1 depletion, while it is accelerated after down-regulation of DNA-PKcs and 53BP1, two DSB repair key factors. In conclusion, the method we describe here can be used to study DSB repair mechanisms, with direct utility for molecularly optimising the knock-out/in outcomes in genome manipulation.

Application of long single-stranded DNA donors in genome editing: generation and validation of mouse mutants

Background

Recent advances in clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) genome editing have led to the use of long single-stranded DNA (lssDNA) molecules for generating conditional mutations. However, there is still limited available data on the efficiency and reliability of this method.

Results

We generated conditional mouse alleles using lssDNA donor templates and performed extensive characterization of the resulting mutations. We observed that the use of lssDNA molecules as donors efficiently yielded founders bearing the conditional allele, with seven out of nine projects giving rise to modified alleles. However, rearranged alleles including nucleotide changes, indels, local rearrangements and additional integrations were also frequently generated by this method. Specifically, we found that alleles containing unexpected point mutations were found in three of the nine projects analyzed. Alleles originating from illegitimate repairs or partial integration of the donor were detected in eight projects. Furthermore, additional integrations of donor molecules were identified in four out of the seven projects analyzed by copy counting. This highlighted the requirement for a thorough allele validation by polymerase chain reaction, sequencing and copy counting of the mice generated through this method. We also demonstrated the feasibility of using lssDNA donors to generate thus far problematic point mutations distant from active CRISPR cutting sites by targeting two distinct genes (Gckr and Rims1). We propose a strategy to perform extensive quality control and validation of both types of mouse models generated using lssDNA donors.

Conclusion

lssDNA donors reproducibly generate conditional alleles and can be used to introduce point mutations away from CRISPR/Cas9 cutting sites in mice. However, our work demonstrates that thorough quality control of new models is essential prior to reliably experimenting with mice generated by this method. These advances in genome editing techniques shift the challenge of mutagenesis from generation to the validation of new mutant models.

Electronic supplementary material

The online version of this article (10.1186/s12915-018-0530-7) contains supplementary material, which is available to authorized users.

Functional genomics and assays of regulatory activity detect mechanisms at loci for lipid traits and coronary artery disease

Many genome-wide association studies (GWAS) have identified signals located in non-coding regions, and an increasing number of functional genomics annotations of regulatory elements and assays of regulatory activity have been used to investigate mechanisms. Genome-wide datasets that characterize chromatin structure help detect potential regulatory elements. Assays to experimentally assess candidate variants include transcriptional reporter assays, and recently, massively parallel reporter assays (MPRAs). Additionally, the effect of candidate regulatory elements and variants on gene expression and function can be evaluated using genomic editing with the CRISPR-Cas9 technology. We highlight some recent studies that employed these strategies to identify variant effects and elucidate molecular and/or biological mechanisms at GWAS loci for lipid traits and coronary artery disease.

Precision genome editing using synthesis-dependent repair of Cas9-induced DNA breaks

Genome editing, the introduction of precise changes in the genome, is revolutionizing our ability to decode the genome. Here we describe a simple method for genome editing in mammalian cells that takes advantage of an efficient mechanism for gene conversion that utilizes linear donors. We demonstrate that PCR fragments containing edits up to 1 kb require only 35-bp homology sequences to initiate repair of Cas9-induced double-stranded breaks in human cells and mouse embryos. We experimentally determine donor DNA design rules that maximize the recovery of edits without cloning or selection.

The RNA-guided DNA endonuclease Cas9 has emerged as a powerful tool for genome engineering. Cas9 creates targeted double-stranded breaks (DSBs) in the genome. Knockin of specific mutations (precision genome editing) requires homology-directed repair (HDR) of the DSB by synthetic donor DNAs containing the desired edits, but HDR has been reported to be variably efficient. Here, we report that linear DNAs (single and double stranded) engage in a high-efficiency HDR mechanism that requires only ∼35 nucleotides of homology with the targeted locus to introduce edits ranging from 1 to 1,000 nucleotides. We demonstrate the utility of linear donors by introducing fluorescent protein tags in human cells and mouse embryos using PCR fragments. We find that repair is local, polarity sensitive, and prone to template switching, characteristics that are consistent with gene conversion by synthesis-dependent strand annealing. Our findings enable rational design of synthetic donor DNAs for efficient genome editing.