Current Protocols in Molecular Biology最新文献

High-throughput cell-based screening assays are valuable tools in the discovery of chemical probes and therapeutic agents. Such assays are designed to examine the effects of small compounds on targets, pathways, or phenotypes participating in normal and disease processes. While most cell-based assays measure single quantities, multiplexed assays seek to address these limitations by obtaining multiple simultaneous measurements. The signals from such measurements should be independently detectable and cover large dynamic ranges. Luciferases are good candidates for generation of such signals. They are genetically encoded, versatile, and cost-effective, and their output signals can be sensitively detected. We recently developed a multiplex luciferase assay that allows monitoring the activity of five experimental pathways against one control simultaneously. We used synthetic assembly cloning to assemble all six luciferase reporter units into a single vector over eight stitching rounds. Because all six reporters are on a single piece of DNA, a single vector ensures stoichiometric ratios of each transcriptional unit in each transfected cell, resulting in lower experimental variation. Our proof-of-concept multiplex hextuple luciferase assay was designed to simultaneously monitor the p53, TGF-β, NF-κβ, c-Myc, and MAPK/JNK signaling pathways. The same synthetic assembly cloning pipeline allows the stitching of numerous other cellular pathway luciferase reporters. Here we present an improved three-step synthetic assembly protocol to quickly and efficiently generate multiplex hextuple luciferase reporter plasmids for other signaling pathways of interest. This improved assembly protocol provides the opportunity to analyze any five desired pathways at once much more quickly. Protocols are provided on how to prepare DNA components and destination vector plasmids, design synthetic DNA, perform assembly cloning of new transcriptional reporter elements, implement multipartite synthetic assembly cloning of single-pathway luciferase reporters, and carry out one-step assembly of final multiplex hextuple luciferase vectors. We present protocols on how to perform multiplex hextuple luciferase in an accompanying Current Protocols in Molecular Biology article. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Preparation of DNA parts and destination vectors for synthetic assembly cloning Basic Protocol 2: DNA synthesis and assembly cloning of a typical transcriptional reporter element Alternate Protocol: DNA synthesis and assembly cloning of a challenging transcriptional reporter element Basic Protocol 3: Multipartite synthetic assembly cloning of individual pathway luciferase reporters Basic Protocol 4: One step assembly into final multiplex hextuple luciferase vectors Support Protocol: Generation of home-made chemocompetent E. coli DH10B-T1R cells.

During the course of their life cycle, most RNAs move between several cellular environments where they associate with different RNA binding proteins (RBPs). Reciprocally, a significant portion of RBPs reside in more than a single cellular compartment, where they can interact with discrete RNAs and even exert distinct biological roles. Proximity-CLIP combines proximity biotinylation of proteins with photoactivatable ribonucleoside-enhanced protein-RNA crosslinking to simultaneously profile the proteome, including RBPs and the RBP-bound transcriptome, in any given subcellular compartment. Here we provide a detailed experimental protocol for Proximity-CLIP along with a simplified non-radioactive, small-RNA cDNA library preparation protocol. Published 2020 U.S. Government. Basic Protocol 1: Cell culture, 4SU labeling, proximity biotinylation, and crosslinking Basic Protocol 2: Cell extraction, streptavidin affinity purification, and on-beads trypsinization Basic Protocol 3: RNA footprints cDNA library preparation Support Protocol: Preparation of RNA-seq libraries from intact RNA.

Methods that enable the construction of recombinant DNA molecules are essential tools for biological research and biotechnology. Golden Gate cloning is used for assembly of multiple DNA fragments in a defined linear order in a recipient vector using a one-pot assembly procedure. Golden Gate cloning is based on the use of a type IIS restriction enzyme for digestion of the DNA fragments and vector. Because restriction sites for the type IIS enzyme used for assembly must be present at the ends of the DNA fragments and vector but absent from all internal sequences, special care must be taken to prepare DNA fragments and the recipient vector with a structure suitable for assembly by Golden Gate cloning. In this article, protocols are presented for preparation of DNA fragments, modules, and vectors suitable for Golden Gate assembly cloning. Additional protocols are presented for assembly of defined parts in a transcription unit, as well as the stitching together of multiple transcription units into multigene constructs by the modular cloning (MoClo) pipeline. © 2020 The Authors.

Basic Protocol 1: Performing a typical Golden Gate cloning reaction

Basic Protocol 2: Accommodating a vector to Golden Gate cloning

Basic Protocol 3: Accommodating an insert to Golden Gate cloning

Basic Protocol 4: Generating small standardized parts compatible with hierarchical modular cloning (MoClo) using level 0 vectors

Alternate Protocol: Generating large standardized parts compatible with hierarchical modular cloning (MoClo) using level –1 vectors

Basic Protocol 5: Assembling transcription units and multigene constructs using level 1, M, and P MoClo vectors

This article describes two methods for amplifying prions present in experimental and clinical samples: the protein misfolding cyclic amplification (PMCA) assay and the real-time quaking-induced conversion (RT-QuIC) assay. Protocols for preparation of amplification substrate and analysis of results are included in addition to those for the individual assays. For each assay, control and suspect samples are mixed with appropriate amplification substrate, which is whole brains from mice in the case of PMCA and recombinant prion protein produced in bacteria for RT-QuIC, followed by cyclic amplification over a number of cycles of sonication (PMCA) or shaking (RT-QuIC) at a consistent incubation temperature. The resultant amplification products are then assessed either by western blotting (PMCA) or based on fluorescent emissions (RT-QuIC). The equipment and expertise necessary for successfully performing either assay vary and will be important factors for individual laboratories to consider when identifying which assay is more appropriate for their experimental design. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Prion amplification via protein misfolding cyclic amplification

Support Protocol 1: Collection of whole brains from mice and preparation of normal brain homogenate

Basic Protocol 2: Prion amplification via real-time quaking-induced conversion

Support Protocol 2: Preparation of recombinant truncated white-tailed-deer prion protein

Somatic liver knockout (SLiK) is a method developed to rapidly generate a liver-specific knockout of one or several genes. This technique combines the strengths of CRISPR/Cas9 gene editing and hydrodynamic tail-vein injection, a simple in vivo method for transfection of hepatocytes, to harness the powerful selection pressure of tyrosinemic livers to replace host hepatocytes with any desired gene deletion. In this protocol, we will describe sgRNA design and cloning, hydrodynamic tail-vein injection of targeting constructs, and screening and validation methods for efficient in vivo gene editing. © 2020 by John Wiley & Sons, Inc.

Support Protocol 1: sgRNA design

Support Protocol 2: sgRNA construction: daisy chaining multiple sgRNAs

Basic Protocol: Delivery of DNA by hydrodynamic tail-vein injection and liver repopulation of edited hepatocytes

Support Protocol 3: Validation of CRISPR/Cas9 cutting in vivo

Many synthetic biologists have adopted methods based on Type IIS restriction enzymes and Golden Gate technology in their cloning procedures, as these enable the combinatorial assembly of modular elements in a very efficient way following standard rules. GoldenBraid (GB) is a Golden Gate–based modular cloning system that, in addition, facilitates the engineering of large multigene constructs and the exchange of DNA parts as result of its iterative cloning scheme. GB was initially developed specifically for plant synthetic biology, and it has been subsequently extended and adapted to other organisms such as Saccharomyces cerevisiae, filamentous fungi, and human cells by incorporating a number of host-specific features into its basic scheme. Here we describe the general GB cloning procedure and provide detailed protocols for its adaptation to filamentous fungi—a GB variant known as FungalBraid. The assembly of a cassette for gene disruption by homologous recombination, a fungal-specific extension of the GB utility, is also shown. Development of FungalBraid was relatively straightforward, as both plants and fungi can be engineered using the same binary plasmids via Agrobacterium-mediated transformation. We also describe the use of a set of web-based tools available at the GB website that assist users in all cloning procedures. The availability of plant and fungal versions of GB will facilitate genetic engineering in these industrially relevant organisms. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Software-assisted modular DNA assembly of a two gene expression-cassette with GB

Basic Protocol 2: Agrobacterium tumefaciens–mediated transformation of filamentous fungi

Basic Protocol 3: Software-assisted modular DNA assembly of a gene disruption-cassette using GB

Basic Protocol 4: Obtaining disruption transformants

MicroRNAs are short non-coding RNAs involved in post-transcriptional gene regulation, and are increasingly considered to be biomarkers for numerous biological processes and human diseases. Current techniques used for microRNA detection can be expensive and labor-intensive, and typically require amplification, labeling, or radioactive probes. In this protocol, we describe a DNA nanoswitch–based microRNA detection assay termed  “miRacles”: microRNA-activated conditional looping of engineered switches. This method uses conformationally responsive DNA nanoswitches that detect the presence of specific microRNAs with a simple and unambiguous gel-shift assay that can be performed on the benchtop. The assay is low cost, minimalistic, and capable of direct detection of specific microRNAs in unprocessed total RNA samples, with no enzymatic amplification, labeling, or special equipment. The protocol for detection of microRNAs in total RNA can be completed in as little as a few hours, making this assay a compelling alternative to qPCR and Northern blotting. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Preparation of DNA nanoswitches

Basic Protocol 2: Detection of microRNAs from total RNA samples

Support Protocol 1: Optional nanoswitch purification by PEG precipitation

Support Protocol 2: Optional nanoswitch purification by liquid chromatography

R-loops are abundant, RNA-containing chromatin structures that form in the genomes of both eukaryotes and prokaryotes. Devising methods to identify the precise genomic locations of R-loops is critical to understand how these structures regulate numerous cellular processes, including replication, termination, and chromosome segregation, and how their unscheduled formation results in disease. Here, we describe a new, highly sensitive, and antibody-independent method, MapR, to profile native R-loops genome wide. MapR takes advantage of the natural specificity of the RNase H enzyme to recognize DNA:RNA hybrids, a defining feature of R-loops, and combines it with a CUT&RUN approach to target, cleave, and release R-loops that can then be sequenced. MapR has low background, is faster than current R-loop detection technologies, and can be performed in any cell type without the need to generate stable cell lines. © 2020 by John Wiley & Sons, Inc.

The CRISPR-Cas9 system makes it possible to cause double-strand breaks in specific regions, inducing repair. In the presence of a donor construct, repair can involve insertion or ‘knock-in’ of an exogenous cassette. One common application of knock-in technology is to generate cell lines expressing fluorescently tagged endogenous proteins. The standard approach relies on production of a donor plasmid with ∼500 to 1000 bp of homology on either side of an insertion cassette that contains the fluorescent protein open reading frame (ORF). We present two alternative methods for knock-in of fluorescent protein ORFs into Cas9-expressing Drosophila S2R+ cultured cells, the single-stranded DNA (ssDNA) Drop-In method and the CRISPaint universal donor method. Both methods eliminate the need to clone a large plasmid donor for each target. We discuss the advantages and limitations of the standard, ssDNA Drop-In, and CRISPaint methods for fluorescent protein tagging in Drosophila cultured cells. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Knock-in into Cas9-positive S2R+ cells using the ssDNA Drop-In approach

Basic Protocol 2: Knock-in into Cas9-positive S2R+ cells by homology-independent insertion of universal donor plasmids that provide mNeonGreen (CRISPaint method)

Support Protocol 1: sgRNA design and cloning

Support Protocol 2: ssDNA donor synthesis

Support Protocol 3: Transfection using Effectene

Support Protocol 4: Electroporation of S2R+-MT::Cas9 Drosophila cells

Support Protocol 5: Single-cell isolation of fluorescent cells using FACS