Double-stranded DNA Donor Research Articles

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish.

The advent of targeted CRISPR-Cas nuclease technologies has revolutionized the ability to perform precise genome editing in both established and emerging model systems. CRISPR-Cas genome editing systems use a synthetic guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to specific genomic DNA loci, where the Cas endonuclease generates a double-strand break. The repair of double-strand breaks by intrinsic error-prone mechanisms leads to insertions and/or deletions, disrupting the locus. Alternatively, the inclusion of double-stranded DNA donors or single-stranded DNA oligonucleotides in this process can elicit the inclusion of precise genome edits ranging from single nucleotide polymorphisms to small immunological tags or even large fluorescent protein constructs. However, a major bottleneck in this procedure can be finding and isolating the desired edit in the germline. This protocol outlines a robust method for screening and isolating germline mutations at specific loci in Danio rerio (zebrafish); however, these principles may be adaptable in any model where in vivo sperm collection is possible.

Fluorescent Tagging of Endogenous Beta-Globin Gene in S-HUDEP2 Cell Model for Live Cell Monitoring of Beta-Globin Expression and Regulation

Introduction: The translation of β-globin (HBB) gene is crucial for adult hemoglobin (HbA) formation, and naturally occurring mutations in HBB cause disorders of erythropoiesis. Therefore, understanding the mechanisms governing HBB regulation and expression has been an active area of research. We aim to develop a cell model to monitor HBB expression in live cells as a tool to gain better understanding of HBB expression, regulation and for the discovery of novel therapeutic targets. Methods: Sickle Human Umbilical Cord Derived Erythroid Progenitor cells (S-HUDEP2) are immortalized CD34+ hematopoietic stem and progenitor cells with homozygous sickle mutation that exclusively produce sickle hemoglobin upon erythroid differentiation. To generate the cell model in S-HUDEP2, we delivered HiFi SpCas9/gRNA targeting the C-terminus of HBB together with a double-stranded DNA donor template encoding self-cleaving peptide, eGFP, and β-globin polyA signal flanking homology arms for simultaneous expression of β-globin and eGFP under the HBB promoter. We created this clonal cell model and confirmed biallelic precise in-frame targeted integration by PCR, Sanger sequencing and digital droplet PCR copy number assay to ensure simultaneous monitoring of both HBB alleles. Results: To validate this model, we delivered HiFi SpCas9/R-66S sgRNA as ribonucleoprotein (RNP) targeting HBB currently in pre-clinical studies for sickle cell anemia (SCA). With RNP delivery, CRISPR/Cas9 double-strand breaks (DSBs) will lead to insertions and deletions (indels); we hypothesized that varying indel frequencies and patterns will result in change in GFP expression. After RNP delivery, we conducted 7 days of in vitro cellular expansion to allow for completion of editing and stabilization of GFP signal followed by 5 days of in vitro cellular differentiation. We identified 3 distinct populations based on GFP expression: GFP-, GFPlow, and GFPhigh. These cells were bulk sorted, genomic DNA was extracted and we analyzed indel patterns for each population through targeted amplicon Next Generation Sequencing (NGS). Each population had a unique indel pattern: confirming that GFP expression is based on genotype. In addition, an increase in GFP mean fluorescent intensity (MFI) was seen after in vitro differentiation with preservation of these distinct groups; indicative of expected increase in HBB transcription. Based on these observations, through use of ddPCR copy number assay, we determined that the creation of CRISPR/Cas9 mediated large deletion (>200bp) is the underlying mechanism leading to GFP loss seen in GFP- andGFPlow populations. Nonsense mutations occurring early in exon 1 of HBB resulting in premature translation-termination codons are resistant to nonsense mediated mRNA decay (NMD), a well-known cellular surveillance mechanism. We hypothesized that RNP induced small frameshift indels in exon 1 allow for NMD escape thus preserving HBB translation and GFP expression while large deletions do not, leading to GFP loss. To further validate this model and define its potential for broad applicability, we confirmed its ability to undergo normal differentiation based on erythroid specific cell markers when compared to untagged S-HUDEP2. We also confirmed our model's capacity to form Hb tetramer with α-chains using Hb native PAGE which allows for identification of intact Hb tetramers based on reference migration. Conclusion: We have successfully established and validated a clonal cell model with GFP tagged β-globin, which can be used to monitor HBB expression in live cells via GFP expression, eliminating the need for cellular fixation and HbA antibody staining. Our model may provide a useful tool for a broad range of applications, including the examination of cellular mechanisms underlying HBB expression and the screening of potential therapeutic targets for hemoglobinopathies and other disorders of erythropoiesis.

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (>1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (>1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

CRISPR/Cas9 mediates efficient site-specific mutagenesis of the odorant receptor co-receptor (Orco) in the malaria vector Anopheles sinensis.

Anopheles sinensis is the most widely distributed mosquito species and is the main transmitter of Plasmodium vivax malaria in China. Most previous research has focused on the mechanistic understanding of biological processes in An. sinensis and novel ways of interrupting malaria transmission. However, the development of functional genomics and genetics-based vector control strategies against An. sinensis remain limited because of insufficient site-specific genome editing tools. We report the first successful application of the CRISPR/Cas9 mediated knock-in for highly efficient, site-specific mutagenesis in An. sinensis. The EGFP marker gene driven by the 3 × P3 promoter was precisely integrated into the odorant receptor co-receptor (Orco) by direct injections of Cas9 protein, double-stranded DNA donor, and Orco-gRNA. We achieved a mutation rate of 3.77%, similar to rates in other mosquito species. Precise knock-in at the intended locus was confirmed by polymerase chain reaction (PCR) amplification and sequencing. The Orco mutation severely impaired mosquito sensitivity to some odors and their ability to locate and discriminate a human host. Orco was confirmed as a key mediator of multiple olfactory-driven behaviors in the An. sinensis life cycle, highlighting the importance of Orco as a key molecular target for malaria control. The results also demonstrated that CRISPR/Cas9 was a simple and highly efficient genome editing technique for An. sinensis and could be used to develop genetic control tools for this vector. © 2022 Society of Chemical Industry.

Rapid, efficient and activation-neutral gene editing of polyclonal primary human resting CD4+ T cells allows complex functional analyses

CD4+ T cells are central mediators of adaptive and innate immune responses and constitute a major reservoir for human immunodeficiency virus (HIV) in vivo. Detailed investigations of resting human CD4+ T cells have been precluded by the absence of efficient approaches for genetic manipulation limiting our understanding of HIV replication and restricting efforts to find a cure. Here we report a method for rapid, efficient, activation-neutral gene editing of resting, polyclonal human CD4+ T cells using optimized cell cultivation and nucleofection conditions of Cas9–guide RNA ribonucleoprotein complexes. Up to six genes, including HIV dependency and restriction factors, were knocked out individually or simultaneously and functionally characterized. Moreover, we demonstrate the knock in of double-stranded DNA donor templates into different endogenous loci, enabling the study of the physiological interplay of cellular and viral components at single-cell resolution. Together, this technique allows improved molecular and functional characterizations of HIV biology and general immune functions in resting CD4+ T cells.

Tagging Proteins with Fluorescent Reporters Using the CRISPR/Cas9 System and Double-Stranded DNA Donors.

Macromolecular complexes govern the majority of biological processes and are of great biomedical relevance as factors that perturb interaction networks underlie a number of diseases, and inhibition of protein-protein interactions is a common strategy in drug discovery. Genome editing technologies enable precise modifications in protein coding genes in mammalian cells, offering the possibility to introduce affinity tags or fluorescent reporters for proteomic or imaging applications in the bona fide cellular context. Here we describe a streamlined procedure which uses the CRISPR/Cas9 system and a double-stranded donor plasmid for efficient generation of homozygous endogenously GFP-tagged human cell lines. Establishing cellular models that preserve native genomic regulation of the target protein is instrumental to investigate protein localization and dynamics using fluorescence imaging but also to affinity purify associated protein complexes using anti-GFP antibodies or nanobodies.

Publisher Correction: An efficient gene knock-in strategy using 5'-modified double-stranded DNA donors with short homology arms.

An amendment to this paper has been published and can be accessed via a link at the top of the paper.

A Universal Surrogate Reporter for Efficient Enrichment of CRISPR/Cas9-Mediated Homology-Directed Repair in Mammalian Cells

CRISPR/Cas9-mediated homology-directed repair (HDR) can be leveraged to precisely engineer mammalian genomes. However, the inherently low efficiency of HDR often hampers to identify the desired modified cells. Here, we developed a novel universal surrogate reporter system that efficiently enriches for genetically modified cells arising from CRISPR/Cas9-induced HDR events (namely, the “HDR-USR” system). This episomally based reporter can be self-cleaved and self-repaired via HDR to create a functional puromycin selection cassette without compromising genome integrity. Co-transfection of the HDR-USR system into host cells and transient puromycin selection efficiently achieves enrichment of HDR-modified cells. We tested the system for precision point mutation at 16 loci in different human cell lines and one locus in two rodent cell lines. This system exhibited dramatic improvements in HDR efficiency at a single locus (up to 20.7-fold) and two loci at once (42% editing efficiency compared to zero in the control), as well as greatly improved knockin efficiency (8.9-fold) and biallelic deletion (35.9-fold) at test loci. Further increases were achieved by co-expression of yeast Rad52 and linear single-/double-stranded DNA donors. Taken together, our HDR-USR system provides a simple, robust and efficient surrogate reporter for the enrichment of CRISPR/Cas9-induced HDR-based precision genome editing across various targeting loci in different cell lines.

An efficient gene knock-in strategy using 5'-modified double-stranded DNA donors with short homology arms.

Here, we report a rapid CRISPR-Cas9-mediated gene knock-in strategy that uses Cas9 ribonucleoprotein and 5'-modified double-stranded DNA donors with 50-base-pair homology arms and achieved unprecedented 65/40% knock-in rates for 0.7/2.5 kilobase inserts, respectively, in human embryonic kidney 293T cells. The identified 5'-end modification led to up to a fivefold increase in gene knock-in rates at various genomic loci in human cancer and stem cells.

Genome Editing with CRISPR-Cas9 in Lactobacillus plantarum Revealed That Editing Outcomes Can Vary Across Strains and Between Methods.

Lactic-acid bacteria such as Lactobacillus plantarum are commonly used for fermenting foods and as probiotics, where increasingly sophisticated genome-editing tools are employed to elucidate and enhance these microbes' beneficial properties. The most advanced tools to date utilize an oligonucleotide or double-stranded DNA donor for recombineering and Cas9 for targeted DNA cleavage. As the associated methods are often developed in isolation for one strain, it remains unclear how different Cas9-based editing methods compare across strains. Here, this work directly compares two methods in different strains of L. plantarum: one utilizing a plasmid-encoded recombineering template and another utilizing an oligonucleotide donor and an inducible DNA recombinase. This comparison reveals one instance in which only the recombineering-template method generates desired edits and another instance in which only the oligo method generates desired edits. It is further found that both methods exhibit highly variable success editing the same site across multiple L. plantarum strains. Finally, failure modes are identified for the recombineering-template method, including a consistent genomic deletion and reversion of a point mutation in the recombineering template. This study therefore highlights surprising differences for Cas9-mediated genome editing between methods and related strains, arguing for the need for multiple, distinct methods when performing CRISPR-based editing in bacteria.

Increased levels of RECQ5 shift DNA repair from canonical to alternative pathways.

RECQ5 (RECQL5) is one of several human helicases that dissociates RAD51–DNA filaments. The gene that encodes RECQ5 is frequently amplified in human tumors, but it is not known whether amplification correlates with increased gene expression, or how increased RECQ5 levels affect DNA repair at nicks and double-strand breaks. Here, we address these questions. We show that RECQ5 gene amplification correlates with increased gene expression in human tumors, by in silico analysis of over 9000 individual tumors representing 32 tumor types in the TCGA dataset. We demonstrate that, at double-strand breaks, increased RECQ5 levels inhibited canonical homology-directed repair (HDR) by double-stranded DNA donors, phenocopying the effect of BRCA deficiency. Conversely, at nicks, increased RECQ5 levels stimulated ‘alternative’ HDR by single-stranded DNA donors, which is normally suppressed by RAD51; this was accompanied by stimulation of mutagenic end-joining. Even modest changes (2-fold) in RECQ5 levels caused significant dysregulation of repair, especially HDR. These results suggest that in some tumors, RECQ5 gene amplification may have profound consequences for genomic instability.