Cas9 Fusion Research Articles

A Nanobody Toolbox for Recognizing Distinct Epitopes on Cas9

Cas9s and fusions of Cas9s have emerged as powerful tools for genetic manipulations. Fusions of Cas9 with other DNA editing enzymes have led to variants capable of single base editing and catalytically dead Cas9s have emerged as tools to specifically target desired regions of a genome. Here we describe the generation of a panel of nanobodies directed against three unique epitopes on Streptococcus pyogenes Cas9. The nanobodies were identified from a nanobody library derived from an alpaca that had been immunized with Cas9. The most potent binders recognize Cas9 and RNA bound Cas9 equally well and do not inhibit Cas9 cleavage of target DNA. These nanobodies bind non-overlapping epitopes as determined by ELISA based epitope binning experiments and mass photometry. We present the sequences of these clones and supporting biochemical data so the broader scientific community can access these reagents.

Engineered transcription-associated Cas9 targeting in eukaryotic cells.

DNA targeting Class 2 CRISPR-Cas effector nucleases, including the well-studied Cas9 proteins, evolved protospacer-adjacent motif (PAM) and guide RNA interactions that sequentially license their binding and cleavage activities at protospacer target sites. Both interactions are nucleic acid sequence specific but function constitutively; thus, they provide intrinsic spatial control over DNA targeting activities but naturally lack temporal control. Here we show that engineered Cas9 fusion proteins which bind to nascent RNAs near a protospacer can facilitate spatiotemporal coupling between transcription and DNA targeting at that protospacer: Transcription-associated Cas9 Targeting (TraCT). Engineered TraCT is enabled in eukaryotic yeast or human cells when suboptimal PAM interactions limit basal activity and when one or more nascent RNA substrates are still tethered to the actively transcribed target DNA in cis. Using yeast, we further show that this phenomenon can be applied for selective editing at one of two identical targets in distinct gene loci, or, in diploid allelic loci that are differentially transcribed. Our work demonstrates that temporal control over Cas9's targeting activity at specific DNA sites may be engineered without modifying Cas9's core domains and guide RNA components or their expression levels. More broadly, it establishes co-transcriptional RNA binding as a cis-acting mechanism that can conditionally stimulate CRISPR-Cas DNA targeting in eukaryotic cells.

Exonuclease editor promotes precision of gene editing in mammalian cells

BackgroundMany efforts have been made to improve the precision of Cas9-mediated gene editing through increasing knock-in efficiency and decreasing byproducts, which proved to be challenging.ResultsHere, we have developed a human exonuclease 1-based genome-editing tool, referred to as exonuclease editor. When compared to Cas9, the exonuclease editor gave rise to increased HDR efficiency, reduced NHEJ repair frequency, and significantly elevated HDR/indel ratio. Robust gene editing precision of exonuclease editor was even superior to the fusion of Cas9 with E1B or DN1S, two previously reported precision-enhancing domains. Notably, exonuclease editor inhibited NHEJ at double strand breaks locally rather than globally, reducing indel frequency without compromising genome integrity. The replacement of Cas9 with single-strand DNA break-creating Cas9 nickase further increased the HDR/indel ratio by 453-fold than the original Cas9. In addition, exonuclease editor resulted in high microhomology-mediated end joining efficiency, allowing accurate and flexible deletion of targeted sequences with extended lengths with the aid of paired sgRNAs. Exonuclease editor was further used for correction of DMD patient-derived induced pluripotent stem cells, where 30.0% of colonies were repaired by HDR versus 11.1% in the control.ConclusionsTherefore, the exonuclease editor system provides a versatile and safe genome editing tool with high precision and holds promise for therapeutic gene correction.

Enhancing Temozolomide (TMZ) chemosensitivity using CRISPR-dCas9-mediated downregulation of O6-methylguanine DNA methyltransferase (MGMT).

Glioblastoma (GBM) stands out as the most prevalent and aggressive intracranial tumor, notorious for its poor prognosis. The current standard-of-care for GBM patients involves surgical resection followed by radiotherapy, combined with concurrent and adjuvant chemotherapy using Temozolomide (TMZ). The effectiveness of TMZ primarily relies on the activity of O6-methylguanine DNA methyltransferase (MGMT), which removes alkyl adducts from the O6 position of guanine at the DNA level, thereby counteracting the toxic effects of TMZ. In this study, we employed fusions of catalytically-inactive Cas9 (dCas9) to DNA methyltransferases (dCas9-DNMT3A) to selectively downregulation MGMT transcription by inducing methylation at MGMT promoter and K-M enhancer. Our findings demonstrate a significant reduction in MGMT expression, leading to intensified TMZ sensitivity in the HEK293T cell line. This study serves as a proof of concept for the utilization of CRISPR-based gene suppression to overcome TMZ resistance and enhance the lethal effect of TMZ in glioblastoma tumor cells.

Application of prime editing system to introduce TP53 R248Q hotspot mutation in acute lymphoblastic leukemia cell line.

In childhood acute lymphoblastic leukemia (ALL), TP53 gene mutation is associated with chemoresistance in a certain population of relapsed cases. To directly verify the association of TP53 gene mutation with chemoresistance of relapsed childhood ALL cases and improve their prognosis, the development of appropriate human leukemia models having TP53 mutation in the intrinsic gene is required. Here, we sought to introduce R248Q hotspot mutation into the intrinsic TP53 gene in an ALL cell line, 697, by applying a prime editing (PE) system, which is a versatile genome editing technology. The PE2 system uses an artificial fusion of nickase Cas9 and reverse-transcriptase to directly place new genetic information into a target site through a reverse transcriptase template in the prime editing guide RNA (pegRNA). Moreover, in the advanced PE3b system, single guide RNA (sgRNA) matching the edited sequence is also introduced to improve editing efficiency. The initially obtained MDM2 inhibitor-resistant PE3b-transfected subline revealed disrupted p53 transactivation activity, reduced p53 target gene expression, and acquired resistance to chemotherapeutic agents and irradiation. Although the majority of the subline acquired the designed R248Q and adjacent silent mutations, the insertion of the palindromic sequence in the scaffold hairpin structure of pegRNA and the overlap of the original genomic DNA sequence were frequently observed. Targeted next-generation sequencing reconfirmed frequent edit errors in both PE2 and PE3b-transfected 697 cells, and it revealed frequent successful edits in HEK293T cells. These observations suggest a requirement for further modification of the PE2 and PE3b systems for accurate editing in leukemic cells.

Efficient scar-free knock-ins of several kilobases in plants by engineered CRISPR-Cas endonucleases

In plants and mammals, non-homologous end-joining is the dominant pathway to repair DNA double-strand breaks, making it challenging to generate knock-in events. In this study, we identified two groups of exonucleases from the herpes virus and the bacteriophage T7 families that conferred an up to 38-fold increase in homology-directed repair frequencies when fused to Cas9/Cas12a in a tobacco mosaic virus-based transient assay in Nicotiana benthamiana. We achieved precise and scar-free insertion of several kilobases of DNA both in transient and stable transformation systems. In Arabidopsis thaliana, fusion of Cas9 to a herpes virus family exonuclease led to 10-fold higher frequencies of knock-ins in the first generation of transformants. In addition, we demonstrated stable and heritable knock-ins in wheat in 1% of the primary transformants. Taken together, our results open perspectives for the routine production of heritable knock-in and gene replacement events in plants.

Engineering self-deliverable ribonucleoproteins for genome editing in the brain

The delivery of CRISPR ribonucleoproteins (RNPs) for genome editing in vitro and in vivo has important advantages over other delivery methods, including reduced off-target and immunogenic effects. However, effective delivery of RNPs remains challenging in certain cell types due to low efficiency and cell toxicity. To address these issues, we engineer self-deliverable RNPs that can promote efficient cellular uptake and carry out robust genome editing without the need for helper materials or biomolecules. Screening of cell-penetrating peptides (CPPs) fused to CRISPR-Cas9 protein identifies potent constructs capable of efficient genome editing of neural progenitor cells. Further engineering of these fusion proteins establishes a C-terminal Cas9 fusion with three copies of A22p, a peptide derived from human semaphorin-3a, that exhibits substantially improved editing efficacy compared to other constructs. We find that self-deliverable Cas9 RNPs generate robust genome edits in clinically relevant genes when injected directly into the mouse striatum. Overall, self-deliverable Cas9 proteins provide a facile and effective platform for genome editing in vitro and in vivo.

A novel Cas9 fusion protein promotes targeted genome editing with reduced mutational burden in primary human cells.

Precise genome editing requires the resolution of nuclease-induced DNA double strand breaks (DSBs) via the homology-directed repair (HDR) pathway. In mammals, this is typically outcompeted by non-homologous end-joining (NHEJ) that can generate potentially genotoxic insertion/deletion mutations at DSB sites. Because of higher efficacy, clinical genome editing has been restricted to imperfect but efficient NHEJ-based approaches. Hence, strategies that promote DSB resolution via HDR are essential to facilitate clinical transition of HDR-based editing strategies and increase safety. Here we describe a novel platform that consists of a Cas9 fused to DNA repair factors to synergistically inhibit NHEJ and favor HDR for precise repairing of Cas-induced DSBs. Compared to canonical CRISPR/Cas9, the increase in error-free editing ranges from 1.5-fold to 7-fold in multiple cell lines and in primary human cells. This novel CRISPR/Cas9 platform accepts clinically relevant repair templates, such as oligodeoxynucleotides (ODNs) and adeno-associated virus (AAV)-based vectors, and has a lower propensity to induce chromosomal translocations as compared to benchmark CRISPR/Cas9. The observed reduced mutational burden, resulting from diminished indel formation at on- and off-target sites, provides a remarkable gain in safety and advocates this novel CRISPR system as an attractive tool for therapeutic applications depending on precision genome editing.

Cas9-Geminin and Cdt1-fused anti-CRISPR protein synergistically increase editing accuracy.

Genome editing with CRISPR-Cas9, particularly for therapeutic purposes, should be accomplished via the homology-directed repair (HDR) pathway, which exhibits greater precision than other pathways. However, one of the issues to be solved is that genome editing efficiency with HDR is generally low. A Streptococcus pyogenes Cas9 (SpyCas9) fusion with human Geminin (Cas9-Gem) reportedly increases HDR efficiency slightly. In contrast, we found that regulation of SpyCas9 activity with an anti-CRISPR protein (AcrIIA4) fused to Chromatin licensing and DNA replication factor 1 (Cdt1) significantly increases HDR efficiency and reduces off-target effects. Here, another anti-CRISPR protein, AcrIIA5, was applied, and the combined use of Cas9-Gem and Anti-CRISPR+Cdt1 showed synergistic enhancement of HDR efficiency. The method may be applicable to various anti-CRISPR/CRISPR-Cas combinations.

Examination of the Functional Relationship between PD-L1 DNA Methylation and mRNA Expression in Non-Small-Cell Lung Cancer

Immunotherapy targeting the interaction between programmed cell death protein 1 (PD-1) and programmed death-ligand 1 (PD-L1) is a treatment option for patients with non-small-cell lung cancer (NSCLC). The expression of PD-L1 by the NSCLC cells determines treatment effectiveness, but the relationship between PD-L1 DNA methylation and expression has not been clearly described. We investigated PD-L1 DNA methylation, mRNA expression, and protein expression in NSCLC cell lines and tumor biopsies. We used clustered regularly interspaced short palindromic repeats-associated protein 9 (CRISPR-Cas9) to modify PD-L1 genetic contexts and endonuclease deficient Cas9 (dCas9) fusions with ten-eleven translocation methylcytosine dioxygenase 1 (TET1) and DNA (cytosine-5)-methyltransferase 3A (DNMT3A) to manipulate PD-L1 DNA methylation. In NSCLC cell lines, we identified specific PD-L1 CpG sites with methylation levels inversely correlated with PD-L1 mRNA expression. However, inducing PD-L1 mRNA expression with interferon-γ did not decrease the methylation level for these CpG sites, and using CRISPR-Cas9, we found that the CpG sites did not directly confer a negative regulation. dCas9-TET1 and dCas9-DNMT3A could induce PD-L1 hypo- and hyper-methylation, respectively, with the latter conferring a decrease in expression showing the functional impact of methylation. In NSCLC biopsies, the inverse correlation between the methylation and expression of PD-L1 was weak. We conclude that there is a regulatory link between PD-L1 DNA methylation and expression. However, since these measures are weakly associated, this study highlights the need for further research before PD-L1 DNA methylation can be implemented as a biomarker and drug target for measures to improve the effectiveness of PD-1/PD-L1 immunotherapy in NSCLC.

Heritable transgene-free genome editing in plants by grafting of wild-type shoots to transgenic donor rootstocks

Generation of stable gene-edited plant lines using clustered regularly interspaced short palindromic repeats (CRISPR)–CRISPR-associated protein 9 (Cas9) requires a lengthy process of outcrossing to eliminate CRISPR–Cas9-associated sequences and produce transgene-free lines. We have addressed this issue by designing fusions of Cas9 and guide RNA transcripts to tRNA-like sequence motifs that move RNAs from transgenic rootstocks to grafted wild-type shoots (scions) and achieve heritable gene editing, as demonstrated in wild-type Arabidopsis thaliana and Brassica rapa. The graft-mobile gene editing system enables the production of transgene-free offspring in one generation without the need for transgene elimination, culture recovery and selection, or use of viral editing vectors. We anticipate that using graft-mobile editing systems for transgene-free plant production may be applied to a wide range of breeding programs and crop plants.

CRISPR-Cas-Guided Mutagenesis of Chromosome and Virulence Plasmid in Shigella flexneri by Cytosine Base Editing.

Shigella is a Gram-negative bacterium that invades the human gut epithelium. The resulting infection, shigellosis, is the deadliest bacterial diarrheal disease. Much of the information about the genes dictating the pathophysiology of Shigella, both on the chromosome and the virulence plasmid, was obtained by classical reverse genetics. However, technical limitations of the prevalent mutagenesis techniques restrict the generation of mutants in a single reaction to a small number, preventing large-scale targeted mutagenesis of Shigella and the subsequent assessment of phenotype. We adopted a CRISPR-Cas-dependent approach, where a nickase Cas9 and cytidine deaminase fusion is guided by single guide RNA (sgRNA) to introduce targeted C→T transitions, resulting in internal stop codons and premature termination of translation. In proof-of-principle experiments using an mCherry fluorescent reporter, we were able to generate loss-of-function mutants in both Escherichia coli and Shigella flexneri with up to 100% efficacy. Using a modified fluctuation assay, we determined that under optimized conditions, the frequency of untargeted mutations introduced by the Cas9-deaminase fusion was in the same range as spontaneous mutations, making our method a safe choice for bacterial mutagenesis. Furthermore, we programmed the method to mutate well-characterized chromosomal and plasmid-borne Shigella flexneri genes and found the mutant phenotype to be similar to those of the reported gene deletion mutants, with no apparent polar effects at the phenotype level. This method can be used in a 96-well-plate format to increase the throughput and generate an array of targeted loss-of-function mutants in a few days. IMPORTANCE Loss-of-function mutagenesis is critical in understanding the physiological role of genes. Therefore, high-throughput techniques to generate such mutants are important for facilitating the assessment of gene function at a pace that matches systems biology approaches. However, to our knowledge, no such method was available for generating an array of single gene mutants in an important enteropathogen-Shigella. This pathogen causes high morbidity and mortality in children, and antibiotic-resistant strains are quickly emerging. Therefore, determination of the function of unknown Shigella genes is of the utmost importance to develop effective strategies to control infections. Our present work will bridge this gap by providing a rapid method for generating loss-of-function mutants. The highly effective and specific method has the potential to be programmed to generate multiple mutants in a single, massively parallel reaction. By virtue of plasmid compatibility, this method can be extended to other members of Enterobacteriaceae.

CRISPR Activation/Interference Screen to Identify Genetic Networks in HDAC-Inhibitor-Resistant Cells.

Epigenetic alterations have been identified in various tumor types. In part, these alterations are mediated via increased histone deacetylase activity. Although preclinical results of monotherapies with histone deacetylase inhibitors (HDACi) are promising, success in clinical trials is limited. Reasons for these limitations may be de novo or acquired resistance to HDAC inhibitors that could be overcome with rational combination therapies. This requires knowledge of resistance mechanism along with the involved genetic networks. One way to identify such genetic networks is the implementation of a CRISPR-based technology allowing transcriptional repression (CRISPRi) and activation (CRISPRa) at a genome-wide scale. We describe a simple approach to amplify and validate sgRNA libraries, generate a myeloid progenitor cell line expressing catalytically dead Cas9 (dCas9) fusion proteins with transcriptional effectors to repress or activate genetic regions of interest and demonstrate a complementary genome-wide HDACi resistance screening approach. Furthermore, we present bioinformatics tools for quality control and analysis of the sequencing data.

A Novel CRISPR Interference Effector Enabling Functional Gene Characterization with Synthetic Guide RNAs.

While CRISPR interference (CRISPRi) systems have been widely implemented in pooled lentiviral screening, there has been limited use with synthetic guide RNAs for the complex phenotypic readouts enabled by experiments in arrayed format. Here we describe a novel deactivated Cas9 fusion protein, dCas9-SALL1-SDS3, which produces greater target gene repression than first or second generation CRISPRi systems when used with chemically modified synthetic single guide RNAs (sgRNAs), while exhibiting high target specificity. We show that dCas9-SALL1-SDS3 interacts with key members of the histone deacetylase and Swi-independent three complexes, which are the endogenous functional effectors of SALL1 and SDS3. Synthetic sgRNAs can also be used with in vitro-transcribed dCas9-SALL1-SDS3 mRNA for short-term delivery into primary cells, including human induced pluripotent stem cells and primary T cells. Finally, we used dCas9-SALL1-SDS3 for functional gene characterization of DNA damage host factors, orthogonally to small interfering RNA, demonstrating the ability of the system to be used in arrayed-format screening.

Efficient Genome Editing Achieved via Plug-and-Play Adenovirus Piggyback Transport of Cas9/gRNA Complex on Viral Capsid Surface.

The capacity to efficiently deliver the gene-editing enzyme complex to target cells is favored over other forms of gene delivery as it offers one-time hit-and-run gene editing, thus improving precision and safety and reducing potential immunogenicity against edited cells in clinical applications. Here we performed a proof-of-mechanism study and demonstrated that a simian adenoviral vector for DNA delivery can be repurposed as a robust intracellular delivery platform for a functional Cas9/guide RNA (gRNA) complex to recipient cells. In this system, the clinically relevant adenovirus was genetically engineered with a plug-and-display technology based on SpyTag003/SpyCatcher003 coupling chemistry. Under physiological conditions, an off-the-shelf mixture of viral vector with SpyTag003 incorporated into surface capsid proteins and Cas9 fused with SpyCatcher003 led to a rapid titration reaction yielding adenovirus carrying Cas9SpyCatcher003 on the virus surface. The Cas9 fusion protein-conjugated viruses in the presence of a reporter gRNA delivered gene-editing functions to cells with an efficiency comparable to that of a commercial CRISPR/Cas9 transfection reagent. Our data fully validate the adenoviral “piggyback” approach to deliver an intracellularly acting enzyme cargo and, thus, warrant the prospect of engineering tissue-targeted adenovirus carrying Cas9/gRNA for in vivo gene editing.

Robust, Durable Gene Activation In Vivo via mRNA-Encoded Activators.

Programmable control of gene expression via nuclease-null Cas9 fusion proteins has enabled the engineering of cellular behaviors. Here, both transcriptional and epigenetic gene activation via synthetic mRNA and lipid nanoparticle delivery was demonstrated in vivo. These highly efficient delivery strategies resulted in high levels of activation in multiple tissues. Finally, we demonstrate durable gene activation in vivo via transient delivery of a single dose of a gene activator that combines VP64, p65, and HSF1 with a SWI/SNF chromatin remodeling complex component SS18, representing an important step toward gene-activation-based therapeutics. This induced sustained gene activation could be inhibited via mRNA-encoded AcrIIA4, further improving the safety profile of this approach.

Amount of Cas9 protein introduced into mouse embryos via electroporation affects the genome-editing rate.

Genetically engineered animals can be produced quickly using genome editing technology. A new electroporation technique, technique for animal knockout system by electroporation (TAKE), aids in the production of genome-edited animals by introducing nucleases into intact embryos using electroporation instead of microinjection. It is difficult to confirm nuclease delivery into embryos after electroporation using the conventional TAKE method. We previously reported the successful visualization of fluorescently-labeled tracrRNA in embryos after electroporation Cas9 paired with the crRNA:tracrRNA-ATTO550 duplex. However, the amount of fluorescence signal from labeled tracrRNA in embryos did not correlate with the genome editing rate of the offspring. This study examined the visualization of Cas9 protein in embryos after electroporation and its correlation with the genome editing rate of the offspring using a fluorescent Cas9 fusion protein. The fluorescent Cas9 protein was observed in all embryos that survived following electroporation. We found that the efficiency of Cas9 protein delivery into embryos via electroporation depended on the pulse length. Furthermore, we demonstrated that the amount of fluorescent Cas9 protein detected in the embryos correlated with the genome editing efficiency of the embryos. These data indicate that the TAKE method using fluorescently-labeled nucleases can be used to optimize the delivery conditions and verify nuclease delivery into individual embryos prior to embryo transfer for the efficient production of genome-edited animals.

Application of CHyMErA Cas9-Cas12a combinatorial genome-editing platform for genetic interaction mapping and gene fragment deletion screening.

CRISPR-based forward genetic screening represents a powerful approach for the systematic characterization of gene function. Recent efforts have been directed toward establishing CRISPR-based tools for the programmable delivery of combinatorial genetic perturbations, most of which are mediated by a single nuclease and the expression of structurally identical guide backbones from two promoters. In contrast, we have developed CHyMErA (Cas hybrid for multiplexed editing and screening applications), which is based on the co-expression of Cas9 and Cas12a nucleases in conjunction with a hybrid guide RNA (hgRNA) engineered by the fusion of Cas9 and Cas12a guides and expressed from a single U6 promoter. CHyMErA is suitable for the high-throughput deletion of genetic segments including the excision of individual exons. Furthermore, CHyMErA enables the concomitant targeting of two or more genes and can thus be used for the systematic mapping of genetic interactions in mammalian cells. CHyMErA can also be applied for the perturbation of paralogous gene pairs, thereby allowing the capturing of phenotypic roles that would otherwise be masked because of genetic redundancy. Here, we provide instructions for the cloning of hgRNA screening libraries and individual hgRNA constructs and offer guidelines for designing and performing combinatorial pooled genetic screens using CHyMErA. Starting with the generation of Cas9- and Cas12a-expressing cell lines, CHyMErA screening can be implemented within 15-20 weeks.

A Novel Set of Cas9 Fusion Proteins to Stimulate Homologous Recombination: Cas9-HRs.

CRISPR- Cas9 has revolutionized genetic engineering. However, the inability to control double-strand break (DSB) repair has severely limited both therapeutic and academic applications. Many attempts have been made to control DSB repair choice. However, particularly in the case of larger edits, none have been able to bypass the rate-limiting step of homologous recombination (HR): long-range 5' end resection. Here, we describe a novel set of Cas9 fusions, Cas9-HRs, designed to bypass the rate-limiting step of HR repair by simultaneously coupling initial and long-range end resection. Here, we demonstrate that Cas9-HRs can increase the rate of homology directed repair (HDR) by 2- to 2.5-fold and decrease p53 mediated cellular toxicity by two- to fourfold compared to Cas9 and are functional in multiple mammalian cell lines with minimal apparent editing site bias. These properties should make Cas9-HRs an attractive option for applications demanding increased HDR rates for long inserts and/or reduced p53 pathway activation.

Genome-wide programmable transcriptional memory by CRISPR-based epigenome editing

A general approach for heritably altering gene expression has the potential to enable many discovery and therapeutic efforts. Here, we present CRISPRoff-a programmable epigenetic memory writer consisting of a single dead Cas9 fusion protein that establishes DNA methylation and repressive histone modifications. Transient CRISPRoff expression initiates highly specific DNA methylation and gene repression that is maintained through cell division and differentiation of stem cells to neurons. Pairing CRISPRoff with genome-wide screens and analysis of chromatin marks establishes rules for heritable gene silencing. We identify single guide RNAs (sgRNAs) capable of silencing the large majority of genes including those lacking canonical CpG islands (CGIs) and reveal a wide targeting window extending beyond annotated CGIs. The broad ability of CRISPRoff to initiate heritable gene silencing even outside of CGIs expands the canonical model of methylation-based silencing and enables diverse applications including genome-wide screens, multiplexed cell engineering, enhancer silencing, and mechanistic exploration of epigenetic inheritance.