Current Protocols in Molecular Biology最新文献

Cover: Related to Laughery and Wyrick (https://doi.org/10.1002/cpmb.110), The cover image shows the experimental strategy for simple CRISPR genome editing in yeast. The central image depicts how Cas9-induced DNA breaks can be used to generate marker-free genome edits in yeast by homologous recombination (HR) with a repair template. The outer circle depicts the organization of a Cas9/sgRNA expression vector used for simple CRISPR/Cas9 genome editing in yeast.

High-throughput screens in Drosophila melanogaster cell lines have led to discovery of conserved gene functions related to signal transduction, host-pathogen interactions, ion transport, and more. CRISPR/Cas9 technology has opened the door to new types of large-scale cell-based screens. Whereas array-format screens require liquid handling automation and assay miniaturization, pooled-format screens, in which reagents are introduced at random and in bulk, can be done in a standard lab setting. We provide a detailed protocol for conducting and evaluating genome-wide CRISPR single guide RNA (sgRNA) pooled screens in Drosophila S2R+ cultured cells. Specifically, we provide step-by-step instructions for library design and production, optimization of cytotoxin-based selection assays, genome-scale screening, and data analysis. This type of project takes ∼3 months to complete. Results can be used in follow-up studies performed in vivo in Drosophila, mammalian cells, and/or other systems. © 2019 by John Wiley & Sons, Inc.

Basic Protocol: Pooled-format screening with Cas9-expressing Drosophila S2R+ cells in the presence of cytotoxin

Support Protocol 1: Optimization of cytotoxin concentration for Drosophila cell screening

Support Protocol 2: CRISPR sgRNA library design and production for Drosophila cell screening

Support Protocol 3: Barcode deconvolution and analysis of screening data

CRISPR-Cas9 has emerged as a powerful method for editing the genome in a wide variety of species, since it can generate a specific DNA break when targeted by the Cas9-bound guide RNA. In yeast, Cas9-targeted DNA breaks are used to promote homologous recombination with a mutagenic template DNA, in order to rapidly generate genome edits (e.g., DNA substitutions, insertions, or deletions) encoded in the template DNA. Since repeated Cas9-induced DNA breaks select against unedited cells, Cas9 can be used to generate marker-free genome edits. Here, we describe a simple protocol for constructing Cas9-expressing plasmids containing a user-designed guide RNA, as well as protocols for using these plasmids for efficient genome editing in yeast. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Constructing the guide RNA expression vector

Basic Protocol 2: Preparing double-stranded oligonucleotide repair template

Alternate Protocol 1: Preparing a single-stranded oligonucleotide repair template

Basic Protocol 3: Induce genome editing by co-transformation of yeast

Basic Protocol 4: Screening for edited cells

Basic Protocol 5: Removing sgRNA/CAS9 expression vector

Alternate Protocol 2: Removing pML107-derived sgRNA/CAS9 expression vector

Ribosome profiling quantifies the genome-wide ribosome occupancy of transcripts. With the integration of matched RNA sequencing data, the translation efficiency (TE) of genes can be calculated to reveal translational regulation. This layer of gene-expression regulation is otherwise difficult to assess on a global scale and generally not well understood in the context of human disease. Current statistical methods to calculate differences in TE have low accuracy, cannot accommodate complex experimental designs or confounding factors, and do not categorize genes into buffered, intensified, or exclusively translationally regulated genes. This article outlines a method [referred to as deltaTE (ΔTE), standing for change in TE] to identify translationally regulated genes, which addresses the shortcomings of previous methods. In an extensive benchmarking analysis, ΔTE outperforms all methods tested. Furthermore, applying ΔTE on data from human primary cells allows detection of substantially more translationally regulated genes, providing a clearer understanding of translational regulation in pathogenic processes. In this article, we describe protocols for data preparation, normalization, analysis, and visualization, starting from raw sequencing files. © 2019 The Authors.

Basic Protocol: One-step detection and classification of differential translation efficiency genes using DTEG.R

Alternate Protocol: Step-wise detection and classification of differential translation efficiency genes using R

Support Protocol: Workflow from raw data to read counts

Over the past century, formalin-fixed, paraffin-embedded (FFPE) tissue samples have represented the standard for basic histology and immunostaining. However, FFPE has several limitations and less stringent tissue preservation methods are required for the visualization of nucleic acids at high resolution, particularly those that are expressed at low levels. Here, we describe the FFPE properties that negatively impact RNA integrity, an alternative tissue preservation technique that prevents RNA loss, and the steps necessary to optimize slide preparation for single-molecule RNA fluorescent in situ hybridization (smRNA-FISH) and imaging by confocal microscopy. This strategy retains RNA quality and eliminates formalin-induced artifacts, thereby producing high-resolution, diffraction-limited confocal images of even rare RNA transcripts in tissues. As non-coding RNAs and alternative splicing of gene isoforms continue to emerge as important regulators of human health and disease, a reliable, cost-effective approach is required to examine the expression and localization of RNA targets in patient samples. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Preparing an RNase-free workstation

Support Protocol 1: Diethyl pyrocarbonate water treatment

Support Protocol 2: Removing RNase contamination from glassware

Basic Protocol 2: BE70 tissue fixation and processing

Basic Protocol 3: Cutting slide sections from paraffin blocks

Basic Protocol 4: Specimen pre-treatment

Basic Protocol 5: RNA fluorescent in situ hybridization labeling

Basic Protocol 6: Slide mounting

Basic Protocol 7: Generating deconvolution-capable confocal micrographs

Over the past decade, transcriptomic studies using next-generation sequencing (NGS)–based RNA sequencing (RNA-Seq) have greatly contributed to characterizing biochemical and physiological changes in cells and tissues across organisms and experimental conditions. Critical steps in RNA-Seq include the preparation of the sequencing library from extracted RNA. Currently, a large panoply of RNA-Seq kits are commercially available. In these kits, conversion of RNA into a sequencing library involves multiple steps, which are labor-intensive, and cost per sample for library preparation may limit routine use of RNA-Seq. Here we describe a simple method for RNA-Seq library construction, referred to as RNA Fragmentation and Sequencing (RF-Seq). RF-Seq requires as little as 10 ng of total RNA and facilitates the sequencing of the 3′ end of mRNAs. RF-Seq involves the fragmentation of total RNA followed by reverse transcription in presence of the oligo(dT) primer/template switch oligonucleotide and a sample barcoding/enrichment within a single PCR tube/well. The sample barcoding/enrichment step provides more flexibility for individual sample handling. The use of just twenty orthogonal Illumina TruSeq HT barcoding primers facilitates the preparation of 96 uniquely labeled RF-Seq libraries in a single 96-well PCR plate. Twelve RF-Seq libraries can be prepared within 4 hr, with an approximate cost of $10/sample. We provide an example of using RF-Seq to measure gene expression upon activation of an innate immune pathway using STING activator in human blood cells, highlighting the potential usefulness of the proposed method in routine transcriptomic applications such as high-throughput drug screening and/or preclinical toxicity assays. © 2019 by John Wiley & Sons, Inc.

Basic Protocol: RNA fragmentation and sequencing (RF-Seq): Cost-effective, time-efficient, and high-throughput 3′ mRNA sequencing library construction in a single tube

The CRISPR-Cas (clustered regularly interspaced short palindromic repeats–CRISPR-associated protein) system is being used successfully for efficient and targeted genome editing in various organisms, including the nematode Caenorhabditis elegans. Recent studies have developed a variety of CRISPR-Cas9 approaches to enhance genome engineering via two major DNA double-strand break repair pathways: nonhomologous end joining and homologous recombination. Here, we describe a protocol for Cas9-mediated C. elegans genome editing together with single guide RNA (sgRNA) and repair template cloning (canonical marker-free and cassette selection methods), as well as injection methods required for delivering Cas9, sgRNAs, and repair template DNA into the germline. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Guide RNA preparation

Alternate Protocol 1: sgRNA cloning using fusion PCR

Basic Protocol 2: Preparation of a repair template for homologous recombination

Alternate Protocol 2: Preparation of repair template donors for the cassette selection method

Basic Protocol 3: Injecting animals

Basic Protocol 4: Screening transgenic worms with marker-free method

Alternate Protocol 3: Screening transgenic worms with cassette selection method

Tagging proteins with fluorescent reporters such as green fluorescent protein (GFP) is a powerful method to determine protein localization, especially when proteins are tagged in the endogenous context to preserve native genomic regulation. However, insertion of fluorescent reporters into the genomes of mammalian cells has required the construction of plasmids containing selection markers and/or extended sequences homologous to the site of insertion (homology arms). Here we describe a streamlined protocol that eliminates all cloning steps by taking advantage of the high propensity of linear DNAs to engage in homology-directed repair of DNA breaks induced by the Cas9 RNA-guided endonuclease. The protocol uses PCR amplicons, or synthetic gene fragments, with short homology arms (30-40 bp) to insert fluorescent reporters at specific genomic locations. The linear DNAs are introduced into cells with preassembled Cas9-crRNA-tracrRNA complexes using one of two transfection procedures, nucleofection or lipofection. The protocol can be completed under a week, with efficiencies ranging from 0.5% to 20% of transfected cells depending on the locus targeted. © 2019 The Authors.

Cover: In Gopalakrishnan and Winston (https://doi.org/10.1002/cpmb.103), Schematic showing the different steps involved in construction of DNA libraries for whole-genome sequencing starting from sonicated genomic DNA. The sequences of the different color-coded regions are highlighted in Table 2.

The identification of transcriptional enhancers and the quantitative assessment of enhancer activities is essential to understanding how regulatory information for gene expression is encoded in animal and human genomes. Further, it is key to understanding how sequence variants affect enhancer function. STARR-seq enables the direct and quantitative assessment of enhancer activity for millions of candidate sequences of arbitrary length and origin in parallel, allowing the screening of entire genomes and the establishment of genome-wide enhancer activity maps.

In STARR-seq, the candidate sequences are cloned downstream of the core promoter into a reporter gene's transcription unit (i.e., the 3′ UTR). Candidates that function as active enhancers lead to the transcription of reporter mRNAs that harbor the candidates’ sequences. This direct coupling of enhancer sequence and enhancer activity in cis enables the straightforward and efficient cloning of complex candidate libraries and the assessment of enhancer activities of millions of candidates in parallel by quantifying the reporter mRNAs by deep sequencing. This article describes how to create focused and genome-wide human STARR-seq libraries and how to perform STARR-seq screens in mammalian cells, and also describes a novel STARR-seq variant (UMI-STARR-seq) that allows the accurate counting of reporter mRNAs for STARR-seq libraries of low complexity. © 2019 The Authors.

Basic Protocol 1: STARR-seq plasmid library cloning

Basic Protocol 2: Mammalian STARR-seq screening protocol

Alternate Protocol: UMI-STARR-seq screening protocol—unique molecular identifier integration

Support Protocol: Transfection of human cells using the MaxCyte STX scalable transfection system