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CRISPR/Cas9-Based Efficient Genome Editing in Clostridium ljungdahlii, an Autotrophic Gas-Fermenting Bacterium.
Acetogenic bacteria have the potential to convert single carbon gases (CO and CO2) into a range of bulk chemicals and fuels. Realization of their full potential is being impeded by the absence of effective genetic tools for high throughput genome modification. Here we report the development of a highly efficient CRISPR/Cas9 system for rapid genome editing of Clostridium ljungdahlii, a paradigm for the commercial production of ethanol from synthesis gas. Following the experimental selection of two promoters (Pthl and ParaE) for expression of cas9 and the requisite single guide RNA (sgRNA), the efficiency of system was tested by making precise deletions of four genes, pta, adhE1, ctf and pyrE. Deletion efficiencies were 100%, >75%, 100% and >50%, respectively. The system overcomes the deficiencies of currently available tools (more rapid, no added antibiotic resistance gene, scarless and minimal polar effects) and will find utility in other acetogens, including the pathogen Clostridium difficile.
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Tracking-seq reveals the heterogeneity of off-target effects in CRISPR-Cas9-mediated genome editing.
The continued development of novel genome editors calls for a universal method to analyze their off-target effects. Here we describe a versatile method, called Tracking-seq, for in situ identification of off-target effects that is broadly applicable to common genome-editing tools, including Cas9, base editors and prime editors. Through tracking replication protein A (RPA)-bound single-stranded DNA followed by strand-specific library construction, Tracking-seq requires a low cell input and is suitable for in vitro, ex vivo and in vivo genome editing, providing a sensitive and practical genome-wide approach for off-target detection in various scenarios. We show, using the same guide RNA, that Tracking-seq detects heterogeneity in off-target effects between different editor modalities and between different cell types, underscoring the necessity of direct measurement in the original system.
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A redox switch regulates the assembly and anti-CRISPR activity of AcrIIC1.
Anti-CRISPRs (Acrs) are natural inhibitors of bacteria's CRISPR-Cas systems, and have been developed as a safeguard to reduce the off-target effects of CRISPR gene-editing technology. Acrs can directly bind to CRISPR-Cas complexes and inhibit their activities. However, whether this process is under regulation in diverse eukaryotic cellular environments is poorly understood. In this work, we report the discovery of a redox switch for NmeAcrIIC1, which regulates NmeAcrIIC1's monomer-dimer interconversion and inhibitory activity on Cas9. Further structural studies reveal that a pair of conserved cysteines mediates the formation of inactive NmeAcrIIC1 dimer and directs the redox cycle. The redox switch also applies to the other two AcrIIC1 orthologs. Moreover, by replacing the redox-sensitive cysteines, we generated a robust AcrIIC1 variant that maintains potent inhibitory activity under various redox conditions. Our results reveal a redox-dependent regulation mechanism of Acr, and shed light on the design of superior Acr for CRISPR-Cas systems.
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Characterization of targeted knock-in achieved via tandem paired nicking mediated by CRISPR/Cas9 nickases.
Targeted knock-in of specific DNA sequences using CRISPR/Cas9 is an advanced technology that enables programmed genome alterations including insertions, deletions, and base substitutions exactly as designed. Despite its utility in life sciences and promise for medical and industrial applications, it remains critical to establish a methodology for highly precise and efficient targeted knock-in to facilitate the practical use of this technology. Tandem paired nicking (TPN) is a genome editing methodology leveraging nicking variants of CRISPR/Cas9 nucleases (Cas9 nickases) to create site-specific nicks within the homologous region of the genome and donor DNA. Such nicking configuration promotes precise and efficient targeted knock-in while repressing the formation of unintended insertions and deletions and p53-mediated DNA damage response. In this study, we conducted a detailed characterization of TPN-based targeted knock-in by performing genome editing assays with various nicking configurations modified from TPN. Our results demonstrated that genomic nicks remarkably contribute to TPN-based targeted knock-in, whereas donor nicks play a less critical role. The introduction of additional nicks beyond the standard two-nick configuration did not further improve the efficiency of TPN-based targeted knock-in. Comparison with other Cas9 nickase-based methodologies for targeted knock-in demonstrated largely equivalent knock-in efficiencies achieved by these methodologies. High-throughput long-read sequencing confirmed a lower incidence of undesired insertions and deletions of various lengths by TPN, in comparison with a conventional Cas9 nuclease-based approach. These findings underscore TPN as a methodology for precise and efficient targeted knock-in, and highlight the broad potential of Cas9 nickase-based targeted knock-in for clinical and industrial applications.
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Guide RNAs containing universal bases enable Cas9/Cas12a recognition of polymorphic sequences.
CRISPR/Cas complexes enable precise gene editing in a wide variety of organisms. While the rigid identification of DNA sequences by these systems minimizes the potential for off-target effects, it consequently poses a problem for the recognition of sequences containing naturally occurring polymorphisms. The presence of genetic variance such as single nucleotide polymorphisms (SNPs) in a gene sequence can compromise the on-target activity of CRISPR systems. Thus, when attempting to target multiple variants of a human gene, or evolved variants of a pathogen gene using a single guide RNA, more flexibility is desirable. Here, we demonstrate that Cas9 can tolerate the inclusion of universal bases in individual guide RNAs, enabling simultaneous targeting of polymorphic sequences. Crucially, we find that specificity is selectively degenerate at the site of universal base incorporation, and remains otherwise preserved. We demonstrate the applicability of this technology to targeting multiple naturally occurring human SNPs with individual guide RNAs and to the design of Cas12a/Cpf1-based DETECTR probes capable of identifying multiple evolved variants of the HIV protease gene. Our findings extend the targeting capabilities of CRISPR/Cas systems beyond their canonical spacer sequences and highlight a use of natural and synthetic universal bases.
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Designing and executing prime editing experiments in mammalian cells.
Prime editing (PE) is a precision gene editing technology that enables the programmable installation of substitutions, insertions and deletions in cells and animals without requiring double-strand DNA breaks (DSBs). The mechanism of PE makes it less dependent on cellular replication and endogenous DNA repair than homology-directed repair-based approaches, and its ability to precisely install edits without creating DSBs minimizes indels and other undesired outcomes. The capabilities of PE have also expanded since its original publication. Enhanced PE systems, PE4 and PE5, manipulate DNA repair pathways to increase PE efficiency and reduce indels. Other advances that improve PE efficiency include engineered pegRNAs (epegRNAs), which include a structured RNA motif to stabilize and protect pegRNA 3' ends, and the PEmax architecture, which improves editor expression and nuclear localization. New applications such as twin PE (twinPE) can precisely insert or delete hundreds of base pairs of DNA and can be used in tandem with recombinases to achieve gene-sized (>5 kb) insertions and inversions. Achieving optimal PE requires careful experimental design, and the large number of parameters that influence PE outcomes can be daunting. This protocol describes current best practices for conducting PE and twinPE experiments and describes the design and optimization of pegRNAs. We also offer guidelines for how to select the proper PE system (PE1 to PE5 and twinPE) for a given application. Finally, we provide detailed instructions on how to perform PE in mammalian cells. Compared with other procedures for editing human cells, PE offers greater precision and versatility, and can be completed within 2-4 weeks.
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SETDB1/ATF7IP regulate the precise genome engineering of HUSH-regulated genes.
BACKGROUND: The use of programmable nucleases has transformed genome editing and functional genomics. Clustered regularly interspaced palindromic repeats (CRISPR)/CRISPR-associated 9 (Cas9) was developed such that targeted genomic lesions [usually DNA double-stranded breaks (DSBs)] could be introduced in vivo with ease and precision. In the presence of homology donors, these lesions facilitate high-efficiency precise genome editing (PGE) via homology-directed repair (HDR) pathways. Because DSBs can lead to genomic instability, however, a large amount of effort has been invested in methodologies (e.g., base editors) that only require nicking the chromosomal DNA on one strand. Indeed, we have demonstrated in human cells that oligodeoxynucleotide (ODN)-mediated PGE using nickase variants of Cas9 can proceed by at least two HDR subpathways termed synthesis-dependent strand annealing (SDSA) and single-stranded DNA incorporation (ssDI). Which pathway is utilized is determined by which chromosomal strand (sense or anti-sense/Watson or Crick) is nicked and by the strandedness (sense or anti-sense/Watson or Crick) of the donor ODN. RESULTS: While the mechanism of mammalian SDSA is moderately well understood, that of ssDI is not. To gain genetic insight into ssDI, we carried out a genome-wide CRISPR knockout screen to identify those genes which, when absent, enable increased ssDI. This screen identified the protein lysine methyl transferase (PKMT) Su(var)3-9, enhancer-of-zeste and trithorax (SET) domain bifurcated histone lysine methyltransferase 1 (SETDB1):activating transcription factor 7-interacting protein (ATF7IP) heterodimer and the downstream human silencing hub (HUSH) complex as strong negative regulators of ssDI. Consistent with their well-known biological effects, the negative regulation of ssDI by SETDB1/ATF7IP and HUSH was specific for transgenic reporters and for a HUSH-regulated single-copy gene, but was not observed at other (non-HUSH regulated) single-copy endogenous loci. CONCLUSIONS: In toto, these experiments underscore the profound impact that chromatin modifiers - and by extension, chromatin structure - have on PGE outcomes. Specifically, we have identified SETDB1/ATF7IP and the HUSH complex as major negative regulators of the HDR subpathway, ssDI, when the target is a transgene. These experiments are a proof-of-principle that chromatin can act as a potent barrier to genetic recombination and they strongly support the feasibility of extending similar chromatin modulating strategies to enhance PGE efficiency at endogenous single-copy loci.
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A High-Throughput Strategy for Dissecting Mammalian Genetic Interactions.
Comprehensive delineation of complex cellular networks requires high-throughput interrogation of genetic interactions. To address this challenge, we describe the development of a multiplex combinatorial strategy to assess pairwise genetic interactions using CRISPR-Cas9 genome editing and next-generation sequencing. We characterize the performance of combinatorial genome editing and analysis using different promoter and gRNA designs and identified regions of the chimeric RNA that are compatible with next-generation sequencing preparation and quantification. This approach is an important step towards elucidating genetic networks relevant to human diseases and the development of more efficient Cas9-based therapeutics.
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Escherichia coli vectors having stringently repressible replication origins allow a streamlining of Crispr/Cas9 gene editing.
Readily curable plasmids facilitate the construction of plasmid-free bacterial strains after the plasmid encoded genes are no longer needed. The most popular of these plasmids features a temperature-sensitive (Ts) pSC101 origin of replication which can readily revert during usage and cannot be used to construct Ts mutations in essential genes. Plasmid pAM34 which contains an IPTG-dependent origin of replication largely overcomes this issue but is limited by carrying the most commonly utilized antibiotic selection and replication origin. This study describes the construction of an expanded series of plasmid vectors having replication origins of p15a, RSF1030 or RSF1031 that like pAM34 have IPTG-dependent replication. Surprisingly, these plasmids can be cured in fewer generations than pAM34. Derivatives of pAM34 with alternative antibiotic selection markers were also constructed. The utility of these vectors is demonstrated in the construction of a CRISPR-Cas9 system consisting of an IPTG-dependent Cas9 plasmid and a curable guide RNA plasmid having a streptomycin counterselection marker. This system was successfully demonstrated by construction of point mutations, deletions and insertions in the E. coli genome with a very high efficiency and in a shorter timescale than extant methods. The plasmids themselves were readily cured either together or singly from the resultant strains with minimal effort.
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An engineered prime editor with enhanced editing efficiency in plants.
Prime editing is a versatile genome-editing technology, but it suffers from low editing efficiency. In the present study, we introduce optimized prime editors with substantially improved editing efficiency. We engineered the Moloney-murine leukemia virus reverse transcriptase by removing its ribonuclease H domain and incorporated a viral nucleocapsid protein with nucleic acid chaperone activity. Each modification independently improved prime editing efficiency by ~1.8-3.4-fold in plant cells. When combined in our engineered plant prime editor (ePPE), the two modifications synergistically enhanced the efficiency of base substitutions, deletions and insertions at various endogenous sites by on average 5.8-fold compared with the original PPE in cell culture. No significant increase in byproducts or off-target editing was observed. We used the ePPE to generate rice plants tolerant to sulfonylurea and imidazolinone herbicides, observing an editing frequency of 11.3% compared with 2.1% using PPE. We also combined ePPE with the previously reported dual-prime editing guide (peg) RNAs and engineered pegRNAs to further increase efficiency.
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