Cas9-induced Double-strand Breaks Research Articles

DNA damage drives antigen diversification through mosaic Variant Surface Glycoprotein (VSG) formation in Trypanosoma brucei.

Antigenic variation, using large genomic repertoires of antigen-encoding genes, allows pathogens to evade host antibody. Many pathogens, including the African trypanosome Trypanosoma brucei, extend their antigenic repertoire through genomic diversification. While evidence suggests that T. brucei depends on the generation of new variant surface glycoprotein (VSG) genes to maintain a chronic infection, a lack of experimentally tractable tools for studying this process has obscured its underlying mechanisms. Here, we present a highly sensitive targeted sequencing approach for measuring VSG diversification. Using this method, we demonstrate that a Cas9-induced DNA double-strand break within the VSG coding sequence can induce VSG recombination with patterns identical to those observed during infection. These newly generated VSGs are antigenically distinct from parental clones and thus capable of facilitating immune evasion. Together, these results provide insight into the mechanisms of VSG diversification and an experimental framework for studying the evolution of antigen repertoires in pathogenic microbes.

Linking CRISPR-Cas9 double-strand break profiles to gene editing precision with BreakTag.

Cas9 can cleave DNA in both blunt and staggered configurations, resulting in distinct editing outcomes, but what dictates the type of Cas9 incisions is largely unknown. In this study, we developed BreakTag, a versatile method for profiling Cas9-induced DNA double-strand breaks (DSBs) and identifying the determinants of Cas9 incisions. Overall, we assessed cleavage by SpCas9 at more than 150,000 endogenous on-target and off-target sites targeted by approximately 3,500 single guide RNAs. We found that approximately 35% of SpCas9 DSBs are staggered, and the type of incision is influenced by DNA:gRNA complementarity and the use of engineered Cas9 variants. A machine learning model shows that Cas9 incision is dependent on the protospacer sequence and that human genetic variation impacts the configuration of Cas9 cuts and the DSB repair outcome. Matched datasets of Cas9 and engineered variant incisions with repair outcomes show that Cas9-mediated staggered breaks are linked with precise, templated and predictable single-nucleotide insertions, demonstrating that a scission-based gRNA design can be used to correct clinically relevant pathogenic single-nucleotide deletions.

Microparticle-mediated CRISPR DNA delivery for genome editing in poplar.

The use of CRISPR/Cas9 is currently the method of choice for precise genome engineering in plants, including in the biomass crop poplar. The most commonly used method for delivering CRISPR/Cas9 and its components in poplar is via Agrobacterium-mediated transformation, that besides the desired gene-editing event also results in stable T-DNA integration. Here we explore the delivery of the gene-editing reagents via DNA-coated microparticle bombardment into the model tree Populus tremula x P. alba to evaluate its potential for developing transgene-free, gene-edited trees, as well as its potential for integrating donor DNA at specific target sites. Using an optimized transformation method, which favors the regeneration of plants that transiently express the genes on the delivered donor DNA, we regenerated gene-edited plants that are free of the Cas9 and the antibiotic resistance-encoding transgenes. In addition, we report the frequent integration of donor DNA fragments at the Cas9-induced double-strand break, opening opportunities toward targeted gene insertions.

High-throughput sgRNA testing reveals rules for Cas9 specificity and DNA repair in tomato cells.

CRISPR/Cas9 technology has the potential to significantly enhance plant breeding. To determine the specificity and the mutagenic spectrum of SpCas9 in tomato, we designed 89g(uide) RNAs targeting genes of the tomato MYB transcription factor family with varying predicted specificities. Plasmids encoding sgRNAs and Cas9 were introduced into tomato protoplasts, and target sites as well as 224 predicted off-target sites were screened for the occurrence of mutations using amplicon sequencing. Algorithms for the prediction of efficacy of the sgRNAs had little predictive power in this system. The analysis of mutations suggested predictable identity of single base insertions. Off-target mutations were found for 13 out of 89 sgRNAs and only occurred at positions with one or two mismatches (at 14 and 3 sites, respectively). We found that PAM-proximal mismatches do not preclude low frequency off-target mutations. Off-target mutations were not found at all 138 positions that had three or four mismatches. We compared off-target mutation frequencies obtained with plasmid encoding sgRNAs and Cas9 with those induced by ribonucleoprotein (RNP) transfections. The use of RNPs led to a significant decrease in relative off-target frequencies at 6 out of 17, no significant difference at 9, and an increase at 2 sites. Additionally, we show that off-target sequences with insertions or deletions relative to the sgRNA may be mutated, and should be considered during sgRNA design. Altogether, our data help sgRNA design by providing insight into the Cas9-induced double-strand break repair outcomes and the occurrence of off-target mutations.

Shuffling the yeast genome using CRISPR/Cas9-generated DSBs that target the transposable Ty1 elements.

Although homologous recombination between transposable elements can drive genomic evolution in yeast by facilitating chromosomal rearrangements, the details of the underlying mechanisms are not fully clarified. In the genome of the yeast Saccharomyces cerevisiae, the most common class of transposon is the retrotransposon Ty1. Here, we explored how Cas9-induced double-strand breaks (DSBs) directed to Ty1 elements produce genomic alterations in this yeast species. Following Cas9 induction, we observed a significant elevation of chromosome rearrangements such as deletions, duplications and translocations. In addition, we found elevated rates of mitotic recombination, resulting in loss of heterozygosity. Using Southern analysis coupled with short- and long-read DNA sequencing, we revealed important features of recombination induced in retrotransposons. Almost all of the chromosomal rearrangements reflect the repair of DSBs at Ty1 elements by non-allelic homologous recombination; clustered Ty elements were hotspots for chromosome rearrangements. In contrast, a large proportion (about three-fourths) of the allelic mitotic recombination events have breakpoints in unique sequences. Our analysis suggests that some of the latter events reflect extensive processing of the broken ends produced in the Ty element that extend into unique sequences resulting in break-induced replication. Finally, we found that haploid and diploid strain have different preferences for the pathways used to repair double-stranded DNA breaks. Our findings demonstrate the importance of DNA lesions in retrotransposons in driving genome evolution.

Efficient CRISPR/Cas9-Assisted Knockin of Large DNA Donors by Pronuclear Microinjection During S-Phase in Mouse Zygotes.

In the CRISPR/Cas9-mediated gene cassette knockin (KI) strategy, a gene cassette is integrated into a target locus through a proper DNA repair pathway after the Cas9-induced double-strand DNA breaks; the activation of the DNA repair pathway is known to be correlated with the cell cycle. Recently, we have reported a new KI approach named SPRINT (S-phase pronuclear injection for targeting)-CRISPR, focusing on the correlation between the cell cycle and the KI efficiency in the mouse zygote microinjection. Our results suggest that the CRISPR-mediated KI with a homologous recombination-based donor vector during S-phase enhances the KI efficiency. For SPRINT-CRISPR, the uniformity of the zygotes in the cell cycle is achieved by in vitro fertilization, and the zygotes are cryopreserved until use. These reproductive techniques are necessary for efficient KI. Furthermore, Piezo-assisted microinjection has been successful in improving the survival rate of the injected embryos. In this chapter, we describe the protocols that focus on the zygote preparation and Piezo-assisted microinjection of the SPRINT-CRISPR method.

Therapeutic Gene Editing of HSCs Ex Vivo without in Vitro Culture Avoids Genotoxicity, Simplifies Procedures, and Preserves Efficiency and Stemness

Therapeutic gene editing of HSCs promises to cure inherited blood disorders. However, current ex vivo gene editing protocols may be associated with genotoxic potential and complex in vitro cell manufacturing procedures. Base and prime editing reduce double strand breaks (DSBs) but are generally less efficient and might have unique genotoxic potential (e.g. untargeted deaminase activity). Here we test the hypothesis that therapeutic gene editing with targeted endonucleases disrupting HbF-repressing elements for b-hemoglobinopathies may be made safer, simpler, and at least as effective by focusing the editing on nondividing cells. Genotoxic risks of DSBs appear to be concentrated in cycling cells. For example, Cas9-induced DSBs can result in long deletions at the on-target site that appear to be mainly the result of unidirectional extensive resection selectively active in S/G2 cell cycle phases1. In addition, DSBs must be resolved prior to mitosis to prevent chromosome mis-segregation leading to copy number variation and cytoplasmic micronuclei2,3. We measured the cell cycle state of mobilized PB CD34+ HSPCs immediately after collection, or after 24 or 48 hours of culture with cytokines (SCF, TPO, and FLT3L each at 100 ng/ml). 100% of cells were in G0/G1 at 0 hours, 94.6%±1.5% at 24 hours, and 76.7%±2.6% at 48 hours. We electroporated CD34+ HSPCs with 0, 24, or 48 hours of cytokine culture before performing 3xNLS-HiFi-SpCas9:sgRNA ribonucleoprotein (RNP) electroporation targeting the BCL11A erythroid enhancer. We employed a flow cytometry assay to quantify micronuclei in edited HSPCs. The fraction of micronuclei was increased in cells electroporated with Cas9:sgRNA RNP following 24 or 48 hours of cytokine stimulation but not in unstimulated cells. We performed ddPCR on sorted nuclei and micronuclei using assays specific to distal chromosome 2p (telomeric to the BCL11A cleavage site) and to chr1 as a reference autosome. We found ~9-fold enrichment of chr2p in micronuclei after 24 or 48 hours of pre-stimulation cytokine culture prior to gene editing, but strikingly did not observe any enrichment of chr2p in micronuclei in cells edited without pre-stimulation culture, suggesting that chromosome mis-segregation and unwanted downstream consequences could be avoided by editing quiescent HSCs (Fig. 1A). In addition, we developed ddPCR assays to measure genome editing outcomes after editing the BCL11A enhancer with paired cleavages (at the +58 and +55 enhancers) separated by 3-kb. These assays measure: 1) programmed 3-kb deletions, 2) programmed 3-kb inversions, or 3) short indels or unedited alleles. Any missing signals (genomic copies fewer than found on a reference nontargeted autosome) may represent long deletions or rearrangements. Editing cells without pre-stimulation culture was associated with undetectable missing signals (inferred long deletions and rearrangements) as compared to editing after 24 or 48 hours of pre-stimulation culture. We compared our standard protocol of 24 hours of pre-stimulation cytokine culture and 24 hours of post-electroporation cytokine culture to a protocol in which CD34+ HSPCs were electroporated immediately after thawing and then cryopreserved 15 minutes after electroporation, without any cytokine culture, before infusion to NBSGW mice. After 16 weeks, HbF levels were potently induced to similar levels after gene editing in recipients of HSPCs without and with cytokine culture (Fig. 1B). Overall human engraftment was similar for all groups, as was multilineage repopulation. Gene edit frequencies were similar for recipients of HSPCs without and with cytokine culture. We observed comparable editing and engraftment potency of HSPCs without as compared to with cytokine culture across mobilization type (G-CSF or plerixafor), donor disease status (healthy and SCD), and sgRNA target sequence. Together, these results suggest that editing HSPCs ex vivo without in vitro culture: avoids unintended on-target genotoxicity potential, including production of micronuclei, long deletions, and rearrangements; simplifies protocols, eliminating time, cost, and complexity; and preserves gene editing efficiency and HbF induction and HSC repopulation potency. Editing nondividing cells selectively could be a straightforward modification to improve ex vivo editing as well as to advance future efforts for safe and potent in vivo editing of HSCs. Figure 1View largeDownload PPTFigure 1View largeDownload PPT Close modal

Multi-pathway DNA-repair reporters reveal competition between end-joining, single-strand annealing and homologous recombination at Cas9-induced DNA double-strand breaks

DNA double-strand breaks (DSB) are repaired by multiple distinct pathways, with outcomes ranging from error-free repair to mutagenesis and genomic loss. DSB-repair pathway cross-talk and compensation is incompletely understood, despite its importance for genomic stability, oncogenesis, and genome editing using CRISPR/Cas9. To address this, we constructed and validated three fluorescent Cas9-based reporters, named DSB-Spectrum, that simultaneously quantify the contribution of multiple DNA repair pathways at a DSB. DSB-Spectrum reporters distinguish between DSB-repair by error-free canonical non-homologous end-joining (c-NHEJ) versus homologous recombination (HR; reporter 1), mutagenic repair versus HR (reporter 2), and mutagenic end-joining versus single strand annealing (SSA) versus HR (reporter 3). Using these reporters, we show that inhibiting the c-NHEJ factor DNA-PKcs increases repair by HR, but also substantially increases mutagenic SSA. Our data indicate that SSA-mediated DSB-repair also occurs at endogenous genomic loci, driven by Alu elements or homologous gene regions. Finally, we demonstrate that long-range end-resection factors DNA2 and Exo1 promote SSA and reduce HR, when both pathways compete for the same substrate. These new Cas9-based DSB-Spectrum reporters facilitate the comprehensive analysis of repair pathway crosstalk and DSB-repair outcome.

Target residence of Cas9-sgRNA influences DNA double-strand break repair pathway choices in CRISPR/Cas9 genome editing

BackgroundDue to post-cleavage residence of the Cas9-sgRNA complex at its target, Cas9-induced DNA double-strand breaks (DSBs) have to be exposed to engage DSB repair pathways. Target interaction of Cas9-sgRNA determines its target binding affinity and modulates its post-cleavage target residence duration and exposure of Cas9-induced DSBs. This exposure, via different mechanisms, may initiate variable DNA damage responses, influencing DSB repair pathway choices and contributing to mutational heterogeneity in genome editing. However, this regulation of DSB repair pathway choices is poorly understood.ResultsIn repair of Cas9-induced DSBs, repair pathway choices vary widely at different target sites and classical nonhomologous end joining (c-NHEJ) is not even engaged at some sites. In mouse embryonic stem cells, weakening the target interaction of Cas9-sgRNA promotes bias towards c-NHEJ and increases target dissociation and reduces target residence of Cas9-sgRNAs in vitro. As an important strategy for enhancing homology-directed repair, inactivation of c-NHEJ aggravates off-target activities of Cas9-sgRNA due to its weak interaction with off-target sites. By dislodging Cas9-sgRNA from its cleaved targets, DNA replication alters DSB end configurations and suppresses c-NHEJ in favor of other repair pathways, whereas transcription has little effect on c-NHEJ engagement. Dissociation of Cas9-sgRNA from its cleaved target by DNA replication may generate three-ended DSBs, resulting in palindromic fusion of sister chromatids, a potential source for CRISPR/Cas9-induced on-target chromosomal rearrangements.ConclusionsTarget residence of Cas9-sgRNA modulates DSB repair pathway choices likely through varying dissociation of Cas9-sgRNA from cleaved DNA, thus widening on-target and off-target mutational spectra in CRISPR/Cas9 genome editing.

Characterization of sequence contexts that favor alternative end joining at Cas9-induced double-strand breaks.

Alternative end joining (alt-EJ) mechanisms, such as polymerase theta-mediated end joining, are increasingly recognized as important contributors to inaccurate double-strand break repair. We previously proposed an alt-EJ model whereby short DNA repeats near a double-strand break anneal to form secondary structures that prime limited DNA synthesis. The nascent DNA then pairs with microhomologous sequences on the other break end. This synthesis-dependent microhomology-mediated end joining (SD-MMEJ) explains many of the alt-EJ repair products recovered following I-SceI nuclease cutting in Drosophila. However, sequence-specific factors that influence SD-MMEJ repair remain to be fully characterized. Here, we expand the utility of the SD-MMEJ model through computational analysis of repair products at Cas9-induced double-strand breaks for 1100 different sequence contexts. We find evidence at single nucleotide resolution for sequence characteristics that drive successful SD-MMEJ repair. These include optimal primer repeat length, distance of repeats from the break, flexibility of DNA sequence between primer repeats, and positioning of microhomology templates relative to preferred primer repeats. In addition, we show that DNA polymerase theta is necessary for most SD-MMEJ repair at Cas9 breaks. The analysis described here includes a computational pipeline that can be utilized to characterize preferred mechanisms of alt-EJ repair in any sequence context.

HideRNAs protect against CRISPR-Cas9 re-cutting after successful single base-pair gene editing

Promiscuous activity of the Streptococcus pyogenes DNA nuclease CRISPR-Cas9 can result in destruction of a successfully modified sequence obtained by templated repair of a Cas9-induced DNA double-strand break. To avoid re-cutting, additional target-site-disruptions (TSDs) are often introduced on top of the desired base-pair alteration in order to suppress target recognition. These TSDs may lower the efficiency of introducing the intended mutation and can cause unexpected phenotypes. Alternatively, successfully edited sites can be protected against Cas9 re-cutting activity. This method exploits the finding that Cas9 complexed to trimmed guideRNAs can still tightly bind specific genomic sequences but lacks nuclease activity. We show here that the presence of a guideRNA plus a trimmed guideRNA that matches the successfully mutated sequence, which we call hideRNA, can enhance the recovery of precise single base-pair substitution events tenfold. The benefit of hideRNAs in generating a single point mutation was demonstrated in cell lines using plasmid-based delivery of CRISPR-Cas9 components and in mouse zygotes injected with Cas9/guideRNA plus Cas9/hideRNA ribonucleoprotein complexes. However, hRNA protection sometimes failed, which likely reflects an unfavorable affinity of hRNA/Cas9 versus gRNA/Cas9 for the DNA target site. HideRNAs can easily be implemented into current gene editing protocols and facilitate the recovery of single base-pair substitution. As such, hideRNAs are of great value in gene editing experiments demanding high accuracy.

Investigating CRISPR/Cas9 gene drive for production of disease-preventing prion gene alleles.

Prion diseases are a group of fatal neurodegenerative disorders that includes chronic wasting disease, which affects cervids and is highly transmissible. Given that chronic wasting disease prevalence exceeds 30% in some endemic areas of North America, and that eventual transmission to other mammalian species, potentially including humans, cannot be ruled out, novel control strategies beyond population management via hunting and/or culling must be investigated. Prion diseases depend upon post-translational conversion of the cellular prion protein, encoded by the Prnp gene, into a disease-associated conformation; ablation of cellular prion protein expression, which is generally well-tolerated, eliminates prion disease susceptibility entirely. Inspired by demonstrations of gene drive in caged mosquito species, we aimed to test whether a CRISPR/Cas9-based gene drive mechanism could, in principle, promote the spread of a null Prnp allele among mammalian populations. First, we showed that transient co-expression of Cas9 and Prnp-directed guide RNAs in RK13 cells generates indels within the Prnp open-reading frame, indicating that repair of Cas9-induced double-strand breaks by non-homologous end-joining had taken place. Second, we integrated a ~1.2 kb donor DNA sequence into the Prnp open-reading frame in N2a cells by homology-directed repair following Cas9-induced cleavages and confirmed that integration occurred precisely in most cases. Third, we demonstrated that electroporation of Cas9/guide RNA ribonucleoprotein complexes into fertilised mouse oocytes resulted in pups with a variety of disruptions to the Prnp open reading frame, with a new coisogenic line of Prnp-null mice obtained as part of this work. However, a technical challenge in obtaining expression of Cas9 in the male germline prevented implementation of a complete gene drive mechanism in mice.

CRISPR-Cas9 Targeting of Hepatitis B Virus Covalently Closed Circular DNA Generates Transcriptionally Active Episomal Variants.

ABSTRACTChronic hepatitis B virus (HBV) infection persists due to the lack of therapies that effectively target the HBV covalently closed circular DNA (cccDNA). We used HBV-specific guide RNAs (gRNAs) and CRISPR-Cas9 and determined the fate of cccDNA after gene editing. We set up a ribonucleoprotein (RNP) delivery system in HBV-infected HepG2-NTCP cells. HBV parameters after Cas9 editing were analyzed. Southern blot (SB) analysis and DNA/RNA sequencing (DNA/RNA-seq) were performed to determine the consequences of cccDNA editing and transcriptional activity of mutated cccDNA. Treatment of infected cells with HBV-specific gRNAs showed that CRISPR-Cas9 can efficiently affect HBV replication. The appearance of episomal HBV DNA variants after dual gRNA treatment was observed by PCR, SB analysis, and DNA/RNA-seq. These transcriptionally active variants are the products of simultaneous Cas9-induced double-strand breaks in two target sites, followed by repair and religation of both short and long fragments. Following suppression of HBV DNA replicative intermediates by nucleoside analogs, mutations and formation of smaller transcriptionally active HBV variants were still observed, suggesting that established cccDNA is accessible to CRISPR-Cas9 editing. Targeting HBV DNA with CRISPR-Cas9 leads to cleavage followed by appearance of episomal HBV DNA variants. Effects induced by Cas9 were sustainable after RNP degradation/loss of detection, suggesting permanent changes in the HBV genome instead of transient effects due to transcriptional interference.

A Precise Tool to Edit Human Hematopoietic Stem Cells at High Efficiency and Decipher and Optimize the Different Double Strand Break Repair Outcomes

The CRISPR/Cas9 discovery has revolutionized precise editing of the genome. However, Cas9-induced site-specific double-strand breaks (DSB) are predominantly repaired by the ‘error-prone’ non-homologous end-joining (NHEJ)/alternate NHEJ (aNHEJ) mechanisms, with relatively low precise correction using homology-directed repair (HDR). The HDR-corrected cells can be selected with drugs/surface reporter in cell lines, induced pluripotent cells, mouse embryonic cells. However, selection of gene edited/corrected hematopoietic stem cells (HSC) is not feasible in vitro. HSC remain the most promising targets because of their unique self-renewing, expansion and differentiation potential. Assessment and optimization of editing/correction of the rare human HSC comprising 1-2% of the CD34+ hematopoietic stem and progenitor cell (HSPC) population is particularly challenging, as their selection before/after often results in insufficient cell numbers for detailed molecular analysis; and selection post-editing for clinical translation can reduce recovery/engraftment potential from increased processing. Currently, the readout of the edited HSC is by molecular analysis for NHEJ- and aNHEJ-induced insertions/deletions and HDR detection of total CD34+ cells and assessment of long term repopulating potential (LTRP) in in vivo models. Herein, we report a unique platform that can precisely quantify NHEJ, aNHEJ, HDR and random integration of the HDR donor template in the human HSC using a flow cytometry. We targeted the CD45 surface reporter (PTPRC), present on all leukocytes and HSPC, to induce a DSB in exon 2 of the PTPRC gene using CRISPR/Cas9, and an in-frame promoter-less GFP containing homology template and a downstream tNGFR expression cassette (GFP/tNGFR HDR donor). HSC and HSPC that repaired Cas9-induced DSB via NHEJ on a single allele in diploid cells lost CD45 expression by flow cytometry. Furthermore, the gRNA/Cas9 induced DSB site was designed to have a 3bp micro-homology so that DSB repaired via aNHEJ resulted in cells express CD45 in frame, but a 6bp deletion detectable by TIDE assay. Provision of the GFP/tNGFR HDR donor repair template resulted in GFP and CD45 expressing, but not tNGFR expressing cells, but if the GFP/tNGFR HDR donor integrated randomly, cells were tNGFR+. Using this system, we optimized editing of hematopoietic progenitor cell lines (K562, Kasumi cells) and EBV transformed B-lymphoblasts to >95% efficiency, and molecular analysis of total and CD45+, and CD45- and CD45+GFP+ cell populations confirmed the precision of the flow cytometry based quantification of NHEJ, aNHEJ. In fact, molecular analysis of the CD45+ cells led to the discovery that K562 cells had a high rate of aNHEJ, while this editing via aNHEJ was absent in LCL- and Kasumi- B lymphoid cells. Inhibition of the aNHEJ pathway with small molecule inhibitors/siRNA resulted in loss of the 6bp aNHEJ-mediated deletion in K562 cells. We also found that random integration in LCL, Kasumi and CD34+ cells was a rare event, but occurred with high frequency in K562 cells. Due to the rapid detection of editing events by flow cytometry, we were able to optimize different editing and cell parameters in both cord blood and mobilized peripheral blood derived CD34+ HSPC and CD34+38-90+RA- HSC populations to achieve high viability and 80-87% editing efficiency (CD45-) in HSPC and HSC, and the FACS-based results were similar to molecular analysis. The edited CD34+ HSPC, when injected into NSG mice resulted in robust human cell engraftment, with 45±10% total engraftment, and 39±16% edited cell engraftment (n=12 mice). Using this platform, we improved HDR frequency to 30% and reduced NHEJ events to 60% in hematopoietic cell lines and similar optimization of HDR in HSPC/HSC is underway. In summary, we have developed a novel platform that allows rapid quantification of gene editing by flow cytometry of human HSC at a single cell and a single allele level, in a single step; with concurrent molecular analysis that identifies different DNA repair pathways utilized by different cell types. This platform allows rapid identification of edited events in the rare HSC population, rather than bulk CD34+ HSPC. Furthermore, ability to sort the edited HSC allows precise determination of the genotoxicity of gene editing, and optimization of DSB repair pathway needed, and optimize editing parameters that result in engraftable edited HSC with LTRP. DisclosuresNo relevant conflicts of interest to declare.

Genome-wide interrogation of gene functions through base editor screens empowered by barcoded sgRNAs.

Canonical CRISPR-knockout (KO) screens rely on Cas9-induced DNA double-strand breaks (DSBs) to generate targeted gene KOs. These methodologies may yield distorted results because DSB-associated effects are often falsely assumed to be consequences of gene perturbation itself, especially when high copy-number sites are targeted. In the present study, we report a DSB-independent, genome-wide CRISPR screening method, termed iBARed cytosine base editing-mediated gene KO (BARBEKO). This method leverages CRISPR cytosine base editors for genome-scale KO screens by perturbing gene start codons or splice sites, or by introducing premature termination codons. Furthermore, it is integrated with iBAR, a strategy we devised for improving screening quality and efficiency. By constructing such a cell library through lentiviral infection at a high multiplicity of infection (up to 10), we achieved efficient and accurate screening results with substantially reduced starting cells. More importantly, in comparison with Cas9-mediated fitness screens, BARBEKO screens are no longer affected by DNA cleavage-induced cytotoxicity in HeLa-, K562- or DSB-sensitive retinal pigmented epithelial 1 cells. We anticipate that BARBEKO offers a valuable tool to complement the current CRISPR-KO screens in various settings.

Marker-free quantification of repair pathway utilization at Cas9-induced double-strand breaks.

Genome integrity and genome engineering require efficient repair of DNA double-strand breaks (DSBs) by non-homologous end joining (NHEJ), homologous recombination (HR), or alternative end-joining pathways. Here we describe two complementary methods for marker-free quantification of DSB repair pathway utilization at Cas9-targeted chromosomal DSBs in mammalian cells. The first assay features the analysis of amplicon next-generation sequencing data using ScarMapper, an iterative break-associated alignment algorithm to classify individual repair products based on deletion size, microhomology usage, and insertions. The second assay uses repair pathway-specific droplet digital PCR assays (‘PathSig-dPCR’) for absolute quantification of signature DSB repair outcomes. We show that ScarMapper and PathSig-dPCR enable comprehensive assessment of repair pathway utilization in different cell models, after a variety of experimental perturbations. We use these assays to measure the differential impact of DNA end resection on NHEJ, HR and polymerase theta-mediated end joining (TMEJ) repair. These approaches are adaptable to any cellular model system and genomic locus where Cas9-mediated targeting is feasible. Thus, ScarMapper and PathSig-dPCR allow for systematic fate mapping of a targeted DSB with facile and accurate quantification of DSB repair pathway choice at endogenous chromosomal loci.

A molecular Rosetta Stone to decipher the impact of chromatin features on the repair of Cas9-mediated DNA double-strand breaks

Using a barcoded reporter introduced within a thousand different chromatin locations in human cells, (Schep et al., 2021) characterize repair outcomes of Cas9-induced DNA double-strand breaks (DSBs) and the relative use of DSB repair pathways depending on the local chromatin context.

Impact of chromatin context on Cas9-induced DNA double-strand break repair pathway balance

SummaryDNA double-strand break (DSB) repair is mediated by multiple pathways. It is thought that the local chromatin context affects the pathway choice, but the underlying principles are poorly understood. Using a multiplexed reporter assay in combination with Cas9 cutting, we systematically measure the relative activities of three DSB repair pathways as a function of chromatin context in >1,000 genomic locations. This reveals that non-homologous end-joining (NHEJ) is broadly biased toward euchromatin, while the contribution of microhomology-mediated end-joining (MMEJ) is higher in specific heterochromatin contexts. In H3K27me3-marked heterochromatin, inhibition of the H3K27 methyltransferase EZH2 reverts the balance toward NHEJ. Single-stranded template repair (SSTR), often used for precise CRISPR editing, competes with MMEJ and is moderately linked to chromatin context. These results provide insight into the impact of chromatin on DSB repair pathway balance and guidance for the design of Cas9-mediated genome editing experiments.

Target binding and residence: a new determinant of DNA double-strand break repair pathway choice in CRISPR/Cas9 genome editing.

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) is widely used for targeted genomic and epigenomic modifications and imaging in cells and organisms, and holds tremendous promise in clinical applications. The efficiency and accuracy of the technology are partly determined by the target binding affinity and residence time of Cas9-single-guide RNA (sgRNA) at a given site. However, little attention has been paid to the effect of target binding affinity and residence duration on the repair of Cas9-induced DNA double-strand breaks (DSBs). We propose that the choice of DSB repair pathway may be altered by variation in the binding affinity and residence duration of Cas9-sgRNA at the cleaved target, contributing to significantly heterogeneous mutations in CRISPR/Cas9 genome editing. Here, we discuss the effect of Cas9-sgRNA target binding and residence on the choice of DSB repair pathway in CRISPR/Cas9 genome editing, and the opportunity this presents to optimize Cas9-based technology.

A CRISPR-Cas9, Cre-lox, and Flp-FRT Cascade Strategy for the Precise and Efficient Integration of Exogenous DNA into Cellular Genomes.

We describe a protocol for the precise integration of exogenous DNA into user-defined genomic loci in cultured cells. This strategy first introduces a promoter and a lox site to a specific location via a Cas9-induced double-strand break. Second, a gene of interest (GOI) is inserted into the lox site via Cre-lox recombination. Upon correct insertion, a cis-linked antibiotic resistance gene will be expressed from a promoter introduced into the genome in the first step assuring selection for correct integrants. Last, the selection cassette is excised via a Flp-FRT recombination event, leaving a precisely targeted GOI. This method is broadly applicable to any exogenous DNA to be integrated, choice of integration site, and choice of cell type. The most remarkable aspect of this versatile approach, termed "CasPi" (cascaded precise integration), is that it allows for precise genome targeting with large, frequently complex, and repetitive DNA sequences that do not integrate efficiently or at all with current genome targeting methods.