Efficient CRISPR-Cas genome editing in brown algae.

Targeted genome editing of human pluripotent stem cells (hPSCs) is critical for basic and translational research and can be achieved with site-specific endonucleases. Cpf1 (CRISPR from Prevotella and Francisella) is a programmable DNA endonuclease with AT-rich PAM sequences. In this protocol, we describe procedures for using a single vector system to deliver Cpf1 and CRISPR RNA (crRNA) for genome editing in hPSCs. This protocol enables indel formation and homologous recombination–mediated precise editing at multiple loci. With the delivery of Cpf1 and a single U6 promoter-driven guide RNA array composed of an AAVS1-targeting and a MAFB-targeting crRNA array, efficient multiplex genome editing at the AAVS1 (knockin) and MAFB (knockout) loci in hPSCs could be achieved in a single experiment. The edited hPSCs expressed pluripotency markers and could differentiate into neurons in vitro. This system also generated INS reporter hPSCs with a 6 kb cassette knockin at the INS locus. The INS reporter cells can differentiate into β-cells that express tdTomato and luciferase, permitting fluorescence-activated cell sorting of hPSC-β-cells. By targeted screening of potential off-target sequences that are most homologous to crRNA sequences, no off-target mutations were detected in any of the tested sequences. This work provides an efficient and flexible system for precise genome editing in mammalian cells including hPSCs with the benefits of less off-target effects.Key features• A single-vector system to deliver Cpf1 and crRNA enables the sorting of transfected cells• Efficient and simultaneous multi-modular genome editing exemplified by mutation of MAFB and knockin of AAVS1 loci in a single experiment• Edited PSCs showed minimal off-target effects and can be differentiated into multiple cell types

Protoplast technology enables the identification of efficient multiplex genome editing tools in Phalaenopsis

Fast and efficient CRISPR-mediated genome editing in Aureobasidium using Cas9 ribonucleoproteins

The emergence of clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 technology has dramatically advanced how we manipulate the genome. Regarding in vivo experiments, Cas9‐transgenic animals could provide efficient and complex genome editing. However, this potential has not been fully realized partly due to a lack of convenient platforms and limited examples of successful disease modeling. Here, we devised two doxycycline (Dox)‐inducible Cas9 platforms that efficiently enable conditional genome editing at multiple loci in vitro and in vivo. In these platforms, we took advantage of a site‐specific multi‐segment cloning strategy for rapid and easy integration of multiple single guide (sg)RNAs. We found that a platform containing rtTA at the Rosa26 locus and TRE‐Cas9 together with multiple sgRNAs at the Col1a1 locus showed higher efficiency of inducible insertions and deletions (indels) with minimal leaky editing. Using this platform, we succeeded to model Wilms’ tumor and the progression of intestinal adenomas with multiple mutations including an activating mutation with a large genomic deletion. Collectively, the established platform should make complicated disease modeling in the mouse easily attainable, extending the range of in vivo experiments in various biological fields including cancer research.

Genome editing technology represented by CRISPR-Cas9 had been widely used in many biological fields such as gene function analysis, gene therapy, and crop improvement. However, in the face of the complexity of the eukaryotic genome, the CRISPR-Cas9 genome editing tools have shown an unstable editing efficiency with large variability at different target sites. It was important to further improve the editing efficiency of the CRISPR-Cas9 system among the whole genome. In this study, based on the previous single transcription unit genome editing system (STU-SpCas9), using the ubiquitin-associated domain (UBA) to enhance the stability of Cas9 protein, we constructed three Cas9-UBA fusion systems (SpCas9-SD01, SpCas9-SD02, and SpCas9-SD03). Four different target sites of rice OsPDS, OsDEP1 and OsROC5 genes were chosen to evaluate the genome editing efficiency in rice protoplasts and stable transformed rice plants. The results showed that the fusion of UBA domains did not affect the cleavage mode of Cas9 protein, and effectively increase the editing efficiency of STU-SpCas9 at the target sites. This new CRISPR-Cas9-UBA system provided a new strategy and tool for improving the genome editing efficiency of CRISPR-Cas9 in plants.

Efficient genome editing by controlled release of Cas9 ribonucleoprotein in plant cytosol using polymer-modified microneedle array

The industrial yeast Komagataella phaffii is a highly effective platform for heterologous protein production, owing to its high protein expression and secretion capacity. Heterologous genes and proteins are involved in multiple processes, including transcription, translation, protein folding, modification, transportation, and degradation; however, engineering these proteins and genes is challenging due to inefficient genome editing techniques. We employed Pseudomonas aeruginosa phage single-stranded DNA-annealing protein (SSAP) PapRecT and P. aeruginosa single-stranded DNA-binding protein (SSB) PaSSB to introduce SSAP-SSB-based homology recombination, which facilitated K. phaffii CRISPR-based genome engineering. Specifically, a host-independent method was developed by expressing sgRNA with PapRecT-PaSSB in a single plasmid, with which only a 50 bp short homologous arm (HA) reached a 100% positive rate for CRISPR-based gene insertion, reaching 18 colony-forming units (CFU) per μg of donor DNA. Single deletion using 1000 bp HA attained 100%, reaching 68 CFUs per μg of donor DNA. Using this efficient CRISPR-based genome editing tool, we integrated three genes (INO4, GAL4-like, and PAB1) at three different loci for overexpression to realize the collaborative regulation of human-lactalbumin (α-LA) production. Specifically, we strengthened phospholipid biosynthesis to facilitate endoplasmic reticulum membrane formation and enhanced recombinant protein transcription and translation by overexpressing transcription and translation factors. The final production of α-LA in the 3 L fermentation reached 113.4 mg L-1, two times higher than that of the strain without multiple site gene editing, which is the highest reported titer in K. phaffii. The CRISPR-based genome editing method developed in this study is suitable for the synergistic multiple-site engineering of protein and biochemical biosynthesis pathways to improve the biomanufacturing efficiency.

CRISPR-Cas systems are now widely used for genome editing and transcriptional regulation in diverse organisms. The compact and portable nature of class 2 single effector nucleases, such as Cas9 or Cas12, has facilitated directed genome modifications in plants, animals, and microbes. However, most CRISPR-Cas systems belong to the more prevalent class 1 category, which hinges on multiprotein effector complexes. In the present study, we detail how the native type I-E CRISPR-Cas system, with a 5'-AAA-3' protospacer adjacent motif (PAM) and a 61-nucleotide guide CRISPR RNA (crRNA) can be repurposed for efficient chromosomal targeting and genome editing in Lactobacillus crispatus, an important commensal and beneficial microbe in the vaginal and intestinal tracts. Specifically, we generated diverse mutations encompassing a 643-base pair (bp) deletion (100% efficiency), a stop codon insertion (36%), and a single nucleotide substitution (19%) in the exopolysaccharide priming-glycosyl transferase (p-gtf). Additional genetic targets included a 308-bp deletion (20%) in the prophage DNA packaging Nu1 and a 730-bp insertion of the green fluorescent protein gene downstream of enolase (23%). This approach enables flexible alteration of the formerly genetically recalcitrant species L. crispatus, with potential for probiotic enhancement, biotherapeutic engineering, and mucosal vaccine delivery. These results also provide a framework for repurposing endogenous CRISPR-Cas systems for flexible genome targeting and editing, while expanding the toolbox to include one of the most abundant and diverse systems found in nature.

Genome editing of human induced pluripotent stem cells (hiPSCs) offers unprecedented opportunities for in vitro disease modeling and personalized cell replacement therapy. The introduction of Cas9-directed genome editing has expanded adoption of this approach. However, marker-free genome editing using standard protocols remains inefficient, yielding desired targeted alleles at a rate of ∼1-5%. We developed a protocol based on a doxycycline-inducible Cas9 transgene carried on a piggyBac transposon to enable robust and highly efficient Cas9-directed genome editing, so that a parental line can be expeditiously engineered to harbor many separate mutations. Treatment with doxycycline and transfection with guide RNA (gRNA), donor DNA and piggyBac transposase resulted in efficient, targeted genome editing and concurrent scarless transgene excision. Using this approach, in 7 weeks it is possible to efficiently obtain genome-edited clones with minimal off-target mutagenesis and with indel mutation frequencies of 40-50% and homology-directed repair (HDR) frequencies of 10-20%.

The clustered regularly interspersed short palindromic repeat (CRISPR)/Cas9 system has emerged as an efficient genome engineering method attributed to its high efficiency and versatility. By generating a lethal double-strand DNA break in the target genome, the CRISPR/Cas9 system is capable of selecting the separated crossover events occurring in the traditional genetic manipulation methods in one step, therefore enabling rapid and efficient genome editing in Staphylococcus aureus, including methicillin-resistant S. aureus (MRSA). By engineering the fusion of a cytidine deaminase APOBEC1 and a Cas9 nickase, a base editor was further developed as a highly efficient gene inactivation and point mutation tool in S. aureus. Here we describe a detailed protocol for CRISPR/Cas9-based genome editing in S. aureus, including genome modification and base editing. This protocol outlines in detail the design of primers, the construction and transformation of editing plasmids, as well as the verification of sequence-specific CRISPR/Cas9-mediated mutagenesis in S. aureus.

CRISPR-based DNA editing technologies enable rapid and accessible genome engineering of eukaryotic cells. However, the delivery of genetically encoded CRISPR components remains challenging and sustained Cas9 expression correlates with higher off-target activities, which can be reduced via Cas9-protein delivery. Here we demonstrate that baculovirus, alongside its DNA cargo, can be used to package and deliver proteins to human cells. Using protein-loaded baculovirus (pBV), we demonstrate delivery of Cas9 or base editors proteins, leading to efficient genome and base editing in human cells. By implementing a reversible, chemically inducible heterodimerization system, we show that protein cargoes can selectively and more efficiently be loaded into pBVs (spBVs). Using spBVs we achieved high levels of multiplexed genome editing in a panel of human cell lines. Importantly, spBVs maintain high editing efficiencies in absence of detectable off-targets events. Finally, by exploiting Cas9 protein and template DNA co-delivery, we demonstrate up to 5% site-specific targeted integration of a 1.8 kb heterologous DNA payload using a single spBV in a panel of human cell lines. In summary, we demonstrate that spBVs represent a versatile, efficient and potentially safer alternative for CRISPR applications requiring co-delivery of DNA and protein cargoes.

LNP-mediated delivery of CRISPR RNP for wide-spread in vivo genome editing in mouse cornea.

Target enrichment long-range sequencing (TLES) reveals structural variations missed by PEM-seq

CRISPR-Cas base editing (BE) system is a powerful tool to expand the scope and efficiency of genome editing with single-nucleotide resolution. The editing efficiency, product purity, and off-target effect differ among various BE systems. Herein, we developed CRISPRbase (http://crisprbase.maolab.org), by integrating 1 252 935 records of base editing outcomes in more than 50 cell types from 17 species. CRISPRbase helps to evaluate the putative editing precision of different BE systems by integrating multiple annotations, functional predictions and a blasting system for single-guide RNA sequences. We systematically assessed the editing window, editing efficiency and product purity of various BE systems. Intensive efforts were focused on increasing the editing efficiency and product purity of base editors since the byproduct could be detrimental in certain applications. Remarkably, more than half of cancer-related off-target mutations were non-synonymous and extremely damaging to protein functions in most common tumor types. Luckily, most of these cancer-related mutations were passenger mutations (4840/5703, 84.87%) rather than cancer driver mutations (863/5703, 15.13%), indicating a weak effect of off-target mutations on carcinogenesis. In summary, CRISPRbase is a powerful and convenient tool to study the outcomes of different base editors and help researchers choose appropriate BE designs for functional studies.

DNA double-strand breaks (DSBs) are among the most hazardous cellular damages,potentially leading to cell death or oncogenesis if unrepaired. Genome editing methods,such as the CRISPR/Cas9 system, induce DSBs and utilize these repair pathways for geneknockout and knock-in. Although ionizing radiation also induces DSBs, it is not clearwhether the efficiency of genome editing is affected by ionizing radiation. This studyinvestigated the impact of gamma-ray exposure on the genome editing efficiency of theimproved genome editing via oviductal nucleic acid delivery (i-GONAD)method. Gamma-rays were exposed to pregnant mice receiving i-GONADtargeting the Hr gene, whose mutation causes hair loss in mice. Theexposure on the fertilization day (Day 0) decreased natural delivery rates and littersizes, with notable effects at 0.3 Gy or higher. Although the proportions of hairlessoffspring obtained by i-GONAD differed greatly between single-guide RNAs(sgRNAs) used, total mutation rates, including hairless, mosaic, and indel, wereequivalent. Gamma-ray exposure on Day 0 and the day after fertilization (Day 1) similarlyand almost dose-dependently enhanced the genome editing efficiency evaluated by the totalmutation rate. This study suggests the improvement of genome editing efficiency bygamma-ray exposure, at least in i-GONAD method, potentially facilitatingthe creation of diverse experimental animal models.