The CRISPR Journal

CRISPR/Cas9-mediated transcriptional activation (CRISPRa) is a powerful tool for investigating complex biological phenomena. Although CRISPRa approaches based on VP64 have been widely studied in both cultured cells and in animal models and exhibit great versatility for various cell types and developmental stages in vivo, different dCas9-VP64 versions have not been rigorously compared. Here, we compared different dCas9-VP64 constructs in identical contexts, including the cell lines used and the transfection conditions, for their ability to activate endogenous and exogenous genes. Moreover, we investigated the optimal approach for VP64 addition to VP64- and p300-based constructs. We found that MS2-MCP-scaffolded VP64 enhanced dCas9-VP64 and dCas9-p300 activity better than did direct VP64 fusion to the N-terminus of dCas9. dCas9-VP64+MCP-VP64 and dCas9-p300+MCP-VP64 were superior to VP64-dCas9-VP64 for all target genes tested. Furthermore, multiplexing gRNA expression with dCas9-VP64+MCP-VP64 or dCas9-p300+MCP-VP64 significantly enhanced endogenous gene activation to a level comparable to CRISPRa-SAM with a single gRNA. Our findings demonstrate improvement of the dCas9-VP64 CRISPRa system and contribute to development of a versatile, efficient CRISPRa platform.

We have exploited the repetitive nature of transposable elements of the human genome to generate synthetic circuits. Transposable elements such as LINE-1 and Alu have successfully replicated in mammalian genomes throughout evolution to reach a copy number ranging from thousands to more than a million. Targeting these repetitive elements with programmable DNA nucleases such as CRISPR-Cas9 rapidly induce extremely high levels of cell death. We use this genotoxic feature to build synthetic biocontainment circuits: CRISPR defense system (CRISPR-DS) capable of preventing CRISPR genome editing, and we introduce the proof-of-concept of CRISPR Safety-Switch, an inducible, stringent and non-leaky kill-switch capable of clearing out cell lines resistant to DNA breaks.

The CRISPR/Cas9 system derived from Streptococcus pyogenes (SpCas9) provides unprecedented genome editing capabilities, but the potential for off-target mutations limits its application. In addition to NGG protospacer adjacent motif (PAM), off-target mutations are also associated with noncanonical PAMs, which have not yet been systematically evaluated. Here, we developed a highly sensitive approach that allows systematically analyzing PAM sequences in human cells, and identified multiple alternative PAMs recognized by SpCas9.

Introducing small genetic changes to study specific mutations or reverting clinical mutations to wild type has been an area of interest in precision genomics for several years. In fact, it has been found that nearly 90% of all human pathogenic mutations are caused by small genetic variations, and the methods to efficiently and precisely correct these errors are critically important. One common way to make these small DNA changes is to provide a single stranded DNA (ssDNA) donor containing the desired alteration together with a targeted double-strand break (DSB) at the genomic target. The donor is typically flanked by regions of homology that are often ~30-100bp in length to leverage the homology directed repair (HDR) pathway. Coupling a ssDNA donor with a CRISPR-Cas9 to produce a targeted DSB is one of the most streamlined approaches to introduce small changes. However, in many cell types this approach results in a low rate of incorporation of the desired alteration and has undesired imprecise repair at the 5 or 3 junction due to artifacts of the DNA repair process. We herein report a technology that couples the spatial temporal localization of an ssDNA repair template and leverages the nucleic acid components of the CRISPR-Cas9 system. We show that by direct fusion of an ssDNA template to the trans activating RNA (tracrRNA) to generate an RNA-DNA chimera, termed Donorguide, we recover precise integration of genetic alterations, with both increased integration rates and decreased imprecision at the 5 or 3 junctions relative to an ssODN donor in vitro in HEK293T cells as well as in vivo in zebrafish. Further, we show that this technology can be used to enhance gene conversion with other gene editing tools such as TALENs.

Targeted insertion of exogenous sequences to genomes is useful for therapeutics and biological research. While CRISPR/Cas technologies have been very efficient in gene knockouts by double-strand breaks (DSBs) followed by indel formation through non-homologous end-joining (NHEJ) repair pathway, the precise introduction of new sequences mainly rely on inefficient homology directed repair (HDR) pathways following Cas9-induced DSBs and are restricted to dividing cells. The recent invention of Prime Editing allows short sequences to be precisely inserted at target sites without DSBs. Here, we combine Prime Editing and sequence-specific recombinases and integrases to insert kilobase sequences directionally at target sites. This technique, called PRIMAS for Prime editing, Recombinase, Integrase-mediated Addition of Sequence, will expand our genome editing toolbox for targeted insertion of long sequences up to kilobases and beyond.

Choanoflagellate genetics has undergone rapid and impactful developments in the last decade. Currently, the primary method for genetic modification of choanoflagellates relies on proprietary nucleofection reagents to deliver transgenes for ectopic expression or CRISPR-Cas9 ribonucleoprotein complexes for targeted genome editing. The acquisition of proprietary buffers required for nucleofection can hamper advances in choanoflagellate research due to costs, shipping limitations, and restrictions that prevent buffer components from being optimized for understudied organisms. Therefore, we test whether a low-cost in-house electroporation buffer developed for other systems can replace the proprietary buffer currently used for choanoflagellate transfection. Here, we present an in-house buffer with transfection efficiency comparable to that of the previously established proprietary buffer. This work increases the accessibility of choanoflagellate genetics and can broaden research participation in investigating animal origins.

Synthetic lethality, a genetic interaction involving two or more genes whose combined loss results in cell death, has emerged as a promising strategy for targeted cancer therapy. By exploiting synthetic lethal interactions, cancer cells can be selectively targeted and eradicated while preserving healthy cells, minimizing off-target effects, and reducing toxicity. The development of PARP inhibitors for ovarian and breast cancer patients with BRCA mutations exemplifies the potential of synthetic lethality-based therapy. Various experimental approaches, including CRISPR/Cas9 screens, have been employed to identify synthetic lethal gene pairs. Our lab has developed a CRISPR double knockout library, leveraging the XDeathDB database for candidate gene selection. This comprehensive platform offers insights into 12 cell death modes and 149 cell death hallmark genes. We aim to construct a cell-death double knock-out library using these genes and perform double knock-out screening on MDA-MB-231, a representative cell line for TNBC chemo poor responders. The identified synergistic lethal gene pairs may serve as potential drug targets for treating TNBC.

BackgroundThe widespread application of CRISPR/Cas9 technology has yielded numerous findings in biomedical research in recent years, making it an invaluable tool for gene knockout and for high-throughput screening studies. In (low-throughput) gene knockout studies, editing efficiency is not a major concern because only a few edited clones are necessary for a successful assay. However, in large scale pooled screening studies, editing efficiency is a major concern because each sgRNA has to knockout its target gene in a large cell population in a short period of time. Therefore, a thorough understanding of the role that key factors play in determining CRISPR knockout efficiency is essential to improve the performance of pooled CRISPR screening. MethodsIn this study, cell lines with different expression levels of CAS9 were generated and used to determine gene-editing efficiency. Collections of sgRNAs targeting essential genes were used to study their depletion in the different cell line models. ResultsUsing cell lines with variable expression of Cas9, we confirmed that editing efficiency and speed are mostly dependent on the sgRNA sequence and Cas9 expression, respectively. Importantly, we show that the strategy employed for delivering sgRNAs and Cas9 to cells impacts the performance of high-throughput screens, which is improved in conditions with higher Cas9 expression. ConclusionsOur findings highlight the importance of optimizing Cas9 expression levels when performing gene editing experiments and provide guidance on the necessary decisions for implementing optimal pooled CRISPR screening strategies.

Conditional sex transformation systems are promising tools in the fight against insect pests. In this study, we developed and tested CRISPR-based, tetracycline-repressible sex transformation strains in the Australian sheep blowfly, Lucilia cuprina. Two CRISPR effector molecules, Cas9 and dCas9, were employed to target the sex-determining gene transformer with the goal of turning female blowflies into males. The Cas9 version of the system induced robust knockout of a visual marker gene but failed to trigger sex transformation without external provision of transformer-targeting sgRNAs. Furthermore, we found that dCas9 expression was linked to several deleterious phenotypes, including developmental delays, reduced body weight, and death. Our study provides the first proof-of-concept conditional CRISPR systems in L. cuprina, and suggests that while dCas9 is toxic at high levels in this species, Cas9 is well-tolerated and may be able to induce sex transformation with minor modifications to the system.

The CRISPR-Cas9 system has been widely adopted as a genome editing tool due to its high efficiency and versatility, contributing to the development of various therapeutic strategies. However, its clinical application remains limited by safety concerns, including off-target effects and large-scale chromosomal rearrangements such as translocations and inversions. Recently, the CRISPR-Cas3 system, a Class 1 CRISPR effector complex with unidirectional DNA degradation activity, has gained attention as a potential alternative, offering reduced off-target activity. In this study, we applied the CRISPR-Cas3 system to human T cells and successfully disrupted two clinically relevant genes, T cell receptor alpha constant (TRAC) and beta-2 microglobulin (B2M). These gene deletions were associated with a reduction in both graft-versus-host disease (GVHD) risk and host immune rejection. Importantly, no off-target mutations were detected in CRISPR-Cas3-edited cells, in contrast to the off-target effects observed with CRISPR-Cas9. Furthermore, CAR-T cells generated by deleting TRAC or B2M using CRISPR-Cas3 maintained their antigen-specific cytotoxicity against tumor cells, while exhibiting reduced alloreactivity. These results suggest that CRISPR-Cas3 provides a safer and promising platform for genome editing in T cell engineering, with potential applications in the development of next-generation allogeneic T cell therapies.

Scarless genome editing of induced pluripotent stem cells (iPSCs) is crucial for the precise modeling of genetic disease. Here we present CRISPR Del/Rei, a two-step deletion-reinsertion strategy with high editing efficiency and simple PCR-based screening that generates isogenic clones in ~2 months. We apply our strategy to edit iPSCs at 3 loci with only rare off target editing.

Single-guide RNA lentiviral infection with Cas9 protein electroporation (SLICE) enables CRISPR screening in primary cell types that require transient Cas9 expression yet is limited by scalability and robustness. Here, we introduce dual-guide RNA infection with Cas9 electroporation (DICE), which expresses two guides from the same lentiviral construct that target the same gene. In genome-wide screens, DICE outperformed SLICE in defining essential genes and modulators of PD-L1 expression in IFN gamma activated THP1 cells. Collectively, these data demonstrate that DICE can be utilized for reduced-scale CRISPR screens in cell types with transient Cas9 protein expression without sacrificing screening quality.

The Cas9 nuclease has become central to modern methods and technologies in synthetic biology, largely due to the ease in which it can be targeted to specific DNA loci via guide RNAs (gRNAs). Reports vary widely on the actual specificity of this targeting, with some studies observing 60% of gRNAs possessing no activity against the genome, yet an assumption that inactive gRNAs are rare persists in the E. coli community. To resolve these contradictions, we evaluated the activity of nearly 500,000 unique gRNAs in the E. coli K12 MG1655 genome. We show that the overwhelming majority (at least 93%) of unique gRNAs are functional while only 0.3% are nonfunctional.These nonfunctional gRNAs exhibit strong spacer self-interaction, which can be either excluded using a simple design rule or "repaired" during library design. Finally, this work provides the greater microbial synthetic biology community both a set of nearly half a million E. coli gRNAs that have been empirically evaluated in vivo as well as a thoroughly evaluated experimental procedure, complete with appropriate controls for Cas9 activity, for conducting Cas9 assays in E. coli specifically and bacteria more generally. GRAPHICAL ABSTRACT O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=125 SRC="FIGDIR/small/651106v3_ufig1.gif" ALT="Figure 1"> View larger version (34K): org.highwire.dtl.DTLVardef@1038ddcorg.highwire.dtl.DTLVardef@566e37org.highwire.dtl.DTLVardef@5df6b2org.highwire.dtl.DTLVardef@ef453c_HPS_FORMAT_FIGEXP M_FIG C_FIG

Modeling human disease in human stem cells requires precise, scarless editing of single nucleotide variants (SNV) on one or both chromosomes. Here we describe improved conditions for Cas9 RNP editing of SNVs that yield high rates of biallelic homology-directed repair. To recover both heterozygous and homozygous SNV clones, catalytically inactive dCas9was added to moderate high activity Cas9 RNPs. dCas9 can also block re-cutting and damage to SNV alleles engineered with non-overlapping guide RNAs.

CRISPR-mediated gene activation (CRISPRa) is among the most efficient and reliable strategies for mimicking sustained activation of endogenous promoters and their corresponding genes at physiological levels. By leveraging guide-RNA (gRNA) library design, CRISPRa screens can be applied on a whole-genome scale and are compatible with both arrayed and pooled formats, depending on assay requirements. Compared with conventional arrayed CRISPRa libraries that use single or dual gRNAs and often require multiple gRNA candidates per target, a recently developed CRISPRa library (termed T. gonfio) incorporates four tandem gRNAs per lentivector per target, thereby reducing library complexity and representing the smallest arrayed genome-wide CRISPRa library. To streamline genome-wide arrayed CRISPRa screening, this study developed a high-throughput automated workflow using the Biomek i7 Hybrid liquid-handling platform, integrated with multiple peripheral instruments. The workflow comprises three pipelines: lentiviral library transduction, cell library passaging, and assay processing. These pipelines together establish and maintain the transduced cell library for extended screening times. This enables assay processing at desired extended time points and improves the likelihood of identifying phenotypes that require longer time to develop, making the workflow suitable even for rapidly proliferating cell models. In a pilot arrayed screen using a T. gonfio mini-library targeting kinases and phosphatases, activation of the EPHA2 receptor promoter induced a growth reduction phenotype in the HEK293 cell model. This phenotype was recapitulated in a parallel pooled CRISPRa screen using the same mini-library and further validated in a co-culture assay.

To determine how nuclease deactivated Cas9 (dCas9) or sgRNA expression level affects the knockdown efficiency of CRISPRi, K562 cell clones expressing KRAB-dCas9 protein either with the inducible Tet-on system or with the constitutive SFFV promotor were created by lentiviral transduction, and single clones were selected by fluorescence-activated cell sorting (FACS) for further study. Six genes with various expression levels were targeted using lentiviral sgRNA from two libraries in four cell clones with various KRAB-dCas9 expression levels. We determined the knockdown efficiency and the expression level of the dCas9 protein /sgRNA level by flow cytometry. The cell clone with the highest KRAB-dCas9 expression level achieved effective CRISPRi knockdown, and is statistically different from other clones, indicating enough KRAB-dCas9 expression might be a prerequisite for CRISPRi. Utilizing this clone, we modified the expression level of sgRNA by adopting different multiplicity of infection (MOI)in lentiviral transduction and found that the knockdown efficiency was neither affected by the target gene expression level nor does it correlate with KRAB-dCas9 level, which remained relatively constant (CV=2.2%) across knockdown experiments. 74.72%, 72.28%, 39.08% knockdown of mmadhc, rpia, znf148 genes were achieved, and the knockdown efficiency correlated well with the sgRNA expression level. Linear regression modeling of the data revealed that the knockdown efficiency is significantly affected by both KRAB-dCas9 and sgRNA level, and the sgRNA level has a greater impact, based on the standardized coefficient (0.525 for KRAB-dCas9, 0.981 for sgRNA), indicating that sgRNA level is a major factor affecting CRISPRi efficiency.

Pooled genome-wide CRISPR-Cas9 knockout (CRISPR-KO) screening is a powerful approach for discovering new biology and identifying genetic vulnerabilities in cancers. This approach uses the Cas9 nuclease in combination with sgRNA libraries, typically consisting of 4-8 sgRNAs to induce mutations in each target gene. A critical assumption is that the effect of each sgRNA is solely due to Cas9 editing of the target gene. However, libraries can contain sgRNAs that direct Cas9 to multiple locations, thus potentially introducing bias into gene hit lists and leading to flawed biological hypotheses. Here we have developed GuideRefine, a pipeline to detect multi-targeting and off-targeting sgRNAs. GuideRefine outputs a virtual refined sub-library containing only on-target sgRNAs. Using GuideRefine with T2T-CHM13 as the reference genome, we surveyed the Brunello, TKOv3, Yusa, Avana, and Jacquere libraries, finding that ~7.5% to ~16% of sgRNAs are potentially problematic. We confirmed that multi-targeting sgRNAs disproportionately impair cell fitness and that sgRNAs aligning to more than one location with a single mismatch can also reduce fitness, although to a lesser extent. After flagging problematic sgRNAs and creating virtual ''on-target only'' sub-libraries, ~10% to ~16% of genes lose critical representation (< 3 sgRNAs per gene). Intriguingly, a set of 467 genes, characterised by short CDS length and lower PAM site density, have fewer than three sgRNAs in all sub-libraries, suggesting they cannot be well-targeted using current libraries. We anticipate that GuideRefine, together with caution in assessing the effects of problematic sgRNAs, will help prioritise biologically relevant hits.

The large size of CRISPR-Cas enzymes limits their delivery for therapeutic applications. Cas12j nucleases offers hypercompact alternative but show moderate editing efficiency. To overcome this limitation, we identified eight novel Cas12j orthologues (Cas12j-11 to Cas12j-18) from viral metagenomes. All showed low editing activity in mammalian cells. We engineered T5 exonuclease-Cas12j fusions (T5Exo-Cas12j), two of which, T5Exo-Cas12j-12, and -18 exhibited up to 42% editing in HEK293T and 9% in K-562 cells, outperforming wild-type Cas12j counterparts and comparable to LbCas12a. Intriguingly, robust in cellula editing in both HEK293T and K-562 cells was strictly dependent on the presence of 5'-TAC trinucleotides within the target DNA sequence. Furthermore, we fused the Cas12j orthologues with the TadA8e deaminase and developed base editors, termed Be-(d)Cas12j. Among these, Be-(d)Cas12j-13 demonstrated efficient A-to-G base conversion in mammalian cells. This study expands the CRISPR toolbox by characterizing and engineering novel Cas12j orthologues into compact, high-efficiency genome editors.

CRISPR-Cas9 technology is a powerful tool extensively used for genome editing in mouse and many other species. Streptococcus pyogenes Cas9 efficiently cuts both alleles in mouse zygotes leaving many edited embryos without a functional protein that might be needed to sustain development, to survive postnatally or to reproduce, thus complicating its overwhelmingly advantageous use in making gene modifications. About 25% of mouse genes are essential for embryonic development and another 7% are necessary for fertility, thus for these genes it is desirable to maintain a functional allele to establish viable lines from CRISPR-Cas9 edited mouse embryos. However, exclusive monoallelic editing is challenging to achieve with current CRISPR methods. Controlling the activity of Cas9 in genome editing is an ongoing research field focused on developing new methods to curtail its damage caused by excess of on-target and off-target editing. In this study we describe a novel and a simple method, we termed das-CRISPR, for dCas allele sequestration in combination with CRISPR system, that allows monoallelic editing of targeted allele in mouse and in cultured cell lines. This method incorporates the use of a nuclease deficient deadCas9 (dCas9) present at higher levels than an active Cas9, both complexed with the same single guide RNA (sgRNA) sequence. We showed the delivery of the two proteins as ribonucleoprotein complexes (RNP) into mouse zygotes leads to the generation of viable and fertile mice carrying lethal mutations in an essential gene. We found that greater amounts of dCas9 RNPs bind and protect a target site while the lower amount of functional Cas9 RNPs accessed the unoccupied target site resulting in higher frequency of monoallelic gene editing, compared to using just Cas9 alone. We also showed this method can mitigate and control the activity of Cas9 in mouse NIH3T3 cells in culture to achieve monoallelic editing. This method is a versatile approach to controlling excessive Cas9 activity on-target and off-target both in vitro and in vivo.

Accurate prediction of CRISPR-Cas9 guide RNA (gRNA) editing efficiency remains limited, particularly outside human systems, where models trained on exogenous human datasets show poor generalization. We analyzed Cas9 efficiency and repair outcomes using novel endogenous editing data from four human cell types, two tomato cell types, and cells from giant river prawn and black soldier fly. While integrating publicly available predictors via ensemble frameworks improved performance, our analysis revealed hundreds of novel features affecting activity. Crucially, dominant features related to sites competition for gRNA, and local geometric properties varied across systems, highlighting the strong context dependence of Cas9 efficiency and arguing against a universal model. Interestingly, codon usage bias-based features also emerged as informative predictors, as they are proxies for chromatin accessibility. In contrast, trends in repair outcomes remained conserved. This work provides essential resources for more generalizable CRISPR guide design.