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ABSTRACT Gene-editing tools such as CRISPR-Cas9 have created unprecedented opportunities for genetic studies in plants and animals. We designed a course-based undergraduate research experience (CURE) to train introductory biology students in the concepts and implementation of gene-editing technology as well as develop their soft skills in data management and scientific communication. We present two versions of the course that can be implemented with twice-weekly meetings over a 5-week period. In the remote-learning version, students performed homology searches, designed guide RNAs (gRNAs) and primers, and learned the principles of molecular cloning. This version is appropriate when access to laboratory equipment or in-person instruction is limited, such as during closures that have occurred in response to the COVID-19 pandemic. In person, students designed gRNAs, cloned CRISPR-Cas9 constructs, and performed genetic transformation of Arabidopsis thaliana . Students learned how to design effective gRNA pairs targeting their assigned gene with an 86% success rate. Final exams tested students’ ability to apply knowledge of an unfamiliar genome database to characterize gene structure and to properly design gRNAs. Average final exam scores of ∼73% and ∼84% for in-person and remote-learning CUREs, respectively, indicated that students met learning outcomes. The highly parallel nature of the CURE makes it possible to target dozens to hundreds of genes, depending on the number of sections. Applying this approach in a sensitized mutant background enables focused reverse genetic screens for genetic suppressors or enhancers. The course can be adapted readily to other organisms or projects that employ gene editing.more » « less
CRISPR‐Cas9 genome editing technologies have enabled complex genetic manipulations in situ, including large‐scale, pooled screening approaches to probe and uncover mechanistic insights across various biological processes. The RNA‐programmable nature of CRISPR‐Cas9 greatly empowers tiling mutagenesis approaches to elucidate molecular details of protein function, in particular the interrogation of mechanisms of resistance to small molecules, an approach termed CRISPR‐suppressor scanning. In a typical CRISPR‐suppressor scanning experiment, a pooled library of single‐guide RNAs is designed to target across the coding sequence(s) of one or more genes, enabling the Cas9 nuclease to systematically mutate the targeted proteins and generate large numbers of diverse protein variants in situ. This cellular pool of protein variants is then challenged with drug treatment to identify mutations conferring a fitness advantage. Drug‐resistance mutations identified with this approach can not only elucidate drug mechanism of action but also reveal deeper mechanistic insights into protein structure‐function relationships. In this article, we outline the framework for a standard CRISPR‐suppressor scanning experiment. Specifically, we provide instructions for the design and construction of a pooled sgRNA library, execution of a CRISPR‐suppressor scanning screen, and basic computational analysis of the resulting data. © 2022 Wiley Periodicals LLC.
Basic Protocol 1: Design and generation of a pooled sgRNA library Support Protocol 1: sgRNA library design using command‐line CRISPOR Support Protocol 2: Production and titering of pooled sgRNA library lentivirus Basic Protocol 2: Execution and analysis of a CRISPR‐suppressor scanning experiment
CRISPR‐Cas9‐based technologies have revolutionized experimental manipulation of mammalian genomes. However, limitations regarding the delivery and efficacy of these technologies restrict their application in primary cells. This article describes a protocol for penetrant, reproducible, and fast CRISPR‐Cas9 genome editing in cell cultures derived from primary cells. The protocol employs transient nucleofection of ribonucleoprotein complexes composed of chemically synthesized 2′‐
O‐methyl‐3′phosphorothioate‐modified single guide RNAs (sgRNAs) and purified Cas9 protein. It can be used both for targeted insertion‐deletion mutation (indel) formation at up to >90% efficiency (via use of a single sgRNA) and for targeted deletion of genomic regions (via combined use of multiple sgRNAs). This article provides examples of the nucleofection buffer and programs that are optimal for patient‐derived glioblastoma (GBM) stem‐like cells (GSCs) and human neural stem/progenitor cells (NSCs), but the protocol can be readily applied to other primary cell cultures by modifying the nucleofection conditions. In summary, this is a relatively simple method that can be used for highly efficient and fast gene knockout, as well as for targeted genomic deletions, even in hyperdiploid cells such as many cancer stem‐like cells. © 2020 Wiley Periodicals LLC Basic Protocol: Cas9:sgRNA ribonucleoprotein nucleofection for insertion‐deletion (indel) mutation and genomic deletion generation in primary cell cultures
Phytopathogenic bacteria play important roles in plant productivity, and developments in gene editing have potential for enhancing the genetic tools for the identification of critical genes in the pathogenesis process. CRISPR-based genome editing variants have been developed for a wide range of applications in eukaryotes and prokaryotes. However, the unique mechanisms of different hosts restrict the wide adaptation for specific applications. Here, CRISPR-dCas9 (dead Cas9) and nCas9 (Cas9 nickase) deaminase vectors were developed for a broad range of phytopathogenic bacteria. A gene for a dCas9 or nCas9, cytosine deaminase CDA1, and glycosylase inhibitor fusion protein (cytosine base editor, or CBE) was applied to base editing under the control of different promoters. Results showed that the RecA promoter led to nearly 100% modification of the target region. When residing on the broad host range plasmid pHM1, CBERecApis efficient in creating base edits in strains of
Xanthomonas, Pseudomonas, Erwiniaand Agrobacterium. CBE based on nCas9 extended the editing window and produced a significantly higher editing rate in Pseudomonas. Strains with nonsynonymous mutations in test genes displayed expected phenotypes. By multiplexing guide RNA genes, the vectors can modify up to four genes in a single round of editing. Whole-genome sequencing of base-edited isolates of Xanthomonas oryzaepv. oryzaerevealed guide RNA-independent off-target mutations. Further modifications of the CBE, using a CDA1 variant (CBERecAp-A) reduced off-target effects, providing an improved editing tool for a broad group of phytopathogenic bacteria.
null (Ed.)We report the development of post-transcriptional chemical methods that enable control over CRISPR–Cas9 gene editing activity both in in vitro assays and in living cells. We show that an azide-substituted acyl imidazole reagent (NAI-N 3 ) efficiently acylates CRISPR single guide RNAs (sgRNAs) in 20 minutes in buffer. Poly-acylated (“cloaked”) sgRNA was completely inactive in DNA cleavage with Cas9 in vitro , and activity was quantitatively restored after phosphine treatment. Delivery of cloaked sgRNA and Cas9 mRNA into HeLa cells was enabled by the use of charge-altering releasable transporters (CARTs), which outperformed commercial transfection reagents in transfecting sgRNA co-complexed with Cas9 encoding functional mRNA. Genomic DNA cleavage in the cells by CRISPR–Cas9 was efficiently restored after treatment with phosphine to remove the blocking acyl groups. Our results highlight the utility of reversible RNA acylation as a novel method for temporal control of genome-editing function.more » « less