Current protocols in chemical biology最新文献

The emergence of covalent inhibitors and chemoproteomic probes in translational chemical biology research requires the development of robust biophysical and analytical methods to characterize their complex interactions with target biomolecules. Importantly, these methods must efficiently assess target selectivity and accurately discern noncovalent binding from the formation of resultant covalent adducts. One recently reported covalent chemical tool used in tumor immune oncology, covalent immune recruiters (CIRs), increases the proximity of immune cells and cancer cells, promoting immune recognition and response. Herein we describe biolayer interferometry (BLI) biosensor, flow cytometry, and solution fluorescence-based assay approaches to characterize CIR:antibody binding and CIR-antibody covalent-labeling kinetics. BLI technology, akin to surface plasmon resonance, provides the unique opportunity to investigate molecular binding and labeling kinetics both on a solid surface (Basic Protocol 1) and in solution (Alternate Protocol 1). Here, recruitment of mass-containing proteins to the BLI probe via CIR is measured with high sensitivity and is used as a readout of CIR labeling activity. Further, CIR technology is used to label antibodies with a fluorescent handle. In this system, labeling is monitored via SDS-PAGE with a fluorescence gel imager, where increased fluorescence intensity of a sample reflects increased labeling (Basic Protocol 2). Analysis of CIR:antibody target-specific immune activation is demonstrated with a flow cytometry‒based antibody-dependent cellular phagocytosis (ADCP) assay (Basic Protocol 3). This ADCP protocol may be further used to discern CIR:antibody binding from covalent adduct formation (Alternate Protocol 3). For the protocols described, each method may be used to analyze characteristics of any covalent-tagging or antibody-recruiting small molecule or protein-based technology. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Determining “on-probe” reaction kinetics of CIR1/CIR4 via biolayer interferometry with Octet RED96

Alternate Protocol 1: Determining “in-solution” reaction kinetics of prostate-specific membrane antigen targeting CIR (CIR3) via biolayer interferometry with Octet RED96

Basic Protocol 2: Reaction kinetics of covalently labeled antibodies via fluorescence SDS-PAGE

Basic Protocol 3: Small molecule‒directed antibody-dependent cellular phagocytosis on live human cells measured via flow cytometry

Alternate Protocol 2: Kinetic analysis of CIR3:antibody labeling via antibody-dependent cellular phagocytosis on flow cytometry

Support Protocol 1: Activation of U937 monocytes with interferon γ

Support Protocol 2: Labeling streptavidin beads with biotinylated prostate-specific membrane antigen receptor

Drug-induced liver injury is an important cause of non-approval in drug development and the withdrawal of already approved drugs from the market. Screening human hepatic cell lines for toxicity has been used extensively to predict drug-induced liver injury in preclinical drug development. Assessing hepatic-cell health with more diverse markers will increase the value of in vitro assays and help predict the mechanism of toxicity. We describe three live cell-based assays using HepG2 cells to measure cell health parameters indicative of hepatotoxicity. The first assay measures cellular ATP levels using luciferase. The second and third assays are multiparametric high-content screens covering a panel of cell health markers including cell count, mitochondrial membrane potential and structure, nuclear morphology, vacuolar density, and reactive oxygen species and glutathione levels. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Measurement of cellular ATP content

Basic Protocol 2: High-content analysis assay to assess cell count, mitochondrial membrane potential and structure, and reactive oxygen species

Basic Protocol 3: High-content analysis assay to assess nuclear morphology, vacuoles, and glutathione content

Support Protocol 1: Subculturing and maintaining HepG2 cells

Support Protocol 2: Plating HepG2 cell line

Support Protocol 3: Transferring compounds by pin tool

Support Protocol 4: Generating dose-response curves

Cytidine acetyltransferases are an emerging class of nucleic-acid-modifying enzymes responsible for the establishment of N⁴-acetylcytidine (ac4C) in RNA. In contrast to histone acetyltransferases, whose activity is commonly studied by western blotting, relatively few methods exist for quickly assessing the activity of cytidine acetyltransferases from a biological sample of interest or the distribution of ac4C across different RNA species. In this protocol, we describe a method for analysis of cellular cytidine acetyltransferase activity using dot- and immuno-northern-blotting-based detection. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Detection of N⁴-Acetylcytidine in RNA by dot blotting

Basic Protocol 2: Visualizing N⁴-Acetylcytidine Distribution in RNA by northern blotting

Small molecule microarray (SMM) technology has become a powerful tool used in high-throughput screening for target-based drug discovery. One area in which SMMs have found use is the identification of small molecule ligands for RNA. RNAs with unique secondary or tertiary three-dimensional structures are considered to be attractive targets for small molecules. Complex RNA structures can form hydrophobic pockets suitable for small molecule binding, representing an opportunity for developing novel therapeutics. Our lab has previously taken a target-based approach, screening a single target against many small molecules on an SMM platform. Here, we report a screening protocol for SMMs to investigate multiple RNAs simultaneously using multi-color imaging. By introducing a mixture containing different fluorophore-labeled RNAs, the fluorescence signal of each binding event can be observed simultaneously. Thus, the specificity of a hit compound binding to one RNA target over other highly abundant RNAs (such as tRNA or rRNA) can be easily evaluated. © 2020 Wiley Periodicals LLC.

Basic Protocol: RNA screening on SMMs by multi-color imaging

Support Protocol 1: Preparation of SMM slides

Support Protocol 2: Fluorophore labeling of RNA through maleimide chemistry

Reversible addition-fragmentation chain-transfer (RAFT) polymerization is a commonly used polymerization methodology to generate synthetic polymers. The products of RAFT polymerization, i.e., RAFT polymers, have been widely employed in several biologically relevant areas, including drug delivery, biomedical imaging, and tissue engineering. In this article, we summarize a synthetic methodology to display an azide group at the chain end of a RAFT polymer, thus presenting a reactive site on the polymer terminus. This platform enables a click reaction between azide-terminated polymers and alkyne-containing molecules, providing a broadly applicable scaffold for chemical and bioconjugation reactions on RAFT polymers. We also highlight applications of these azide-terminated RAFT polymers in fluorophore labeling and for promoting organelle targeting capability. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: Synthesis of the azide derivatives of chain transfer agent and radical initiator

Basic Protocol 2: Installation of an azide group on the α-end of RAFT polymers

Alternate Protocol: Installation of an azide group on the ω-end of RAFT polymers

Basic Protocol 3: Click reaction between azide-terminated RAFT polymers and alkyne derivatives

Reverse-polarity activity-based protein profiling (RP-ABPP) is a chemical proteomics approach that uses nucleophilic probes amenable to “click” chemistry deployed into living cells in culture to capture, immunoprecipitate, and identify protein-bound electrophiles. RP-ABPP is used to characterize the structure and function of reactive electrophilic post-translational modifications (PTMs) and the proteins harboring them, which may uncover unknown or novel functions. RP-ABPP has demonstrated utility as a versatile method to monitor the metabolic regulation of electrophilic cofactors, using a pyruvoyl cofactor in S-adenosyl-L-methionine decarboxylase (AMD1), and to discover novel types of electrophilic modifications on proteins in human cells, such as the glyoxylyl modification on secernin-3 (SCRN3). These cofactors cannot be predicted by sequence, and therefore this area is relatively undeveloped. RP-ABPP is the only global, unbiased approach to discover such electrophiles. Here, we describe the utility of these experiments and provide a detailed protocol for de novo discovery, quantitation, and global profiling of electrophilic functionality of proteins. © 2020 The Authors.

Basic Protocol 1: Identification and quantification of probe-reactive proteins

Basic Protocol 2: Characterization of the site of probe labeling

Basic Protocol 3: Determination and quantitation of electrophile structure

The reversible oxidation of protein tyrosine phosphatases (PTPs) impairs their ability to dephosphorylate substrates in vivo. This transient inactivation of PTPs occurs as their conserved catalytic cysteine residue reacts with cellular oxidants thereby abolishing the ability of this reactive cysteine to attack the phosphate of the target substrate. Hence, in vivo, the inhibition of specific PTPs in response to regulated and localized rises in cellular oxidants enables phospho-dependent signaling. We present assays that measure the endogenous activity of specific PTPs that become transiently inactivated in cells exposed to growth factors. Here, we describe the methods and highlight the pitfalls to avoid post-lysis oxidation of PTPs in order to assess the inactivation and the reactivation of PTPs targeted by cellular oxidants in signal transduction. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Cell transfection (optional) Support Protocol: Preparation of degassed lysis buffers Basic Protocol 2: Cellular extraction in anaerobic conditions Basic Protocol 3: Enrichment and activity assay of specific PTPs Alternate Protocol: Measurement of active PTPs via direct cysteinyl labeling.

Metabolomic studies allow a deeper understanding of the processes of a given ecological community than nucleic acid-based surveys alone. In the case of the gut microbiota, a metabolic profile of, for example, a fecal sample provides details about the function and interactions within the distal region of the gastrointestinal tract, and such a profile can be generated in a number of different ways. This unit elaborates on the use of 1D ¹ H NMR spectroscopy as a commonly used method to characterize small-molecule metabolites of the fecal metabonome (meta-metabolome). We describe a set of protocols for the preparation of fecal water extraction, storage, scanning, measurement of pH, and spectral processing and analysis. We also compare the effects of various sample storage conditions for processed and unprocessed samples to provide a framework for comprehensive analysis of small molecules from stool-derived samples. © 2020 Wiley Periodicals LLC Basic Protocol 1: Extracting fecal water from crude fecal samples Alternate Protocol 1: Extracting fecal water from small crude fecal samples Basic Protocol 2: Acquiring NMR spectra of metabolite samples Alternate Protocol 2: Acquiring NMR spectra of metabolite samples using Bruker spectrometer running TopSpin 3.x Alternate Protocol 3: Acquiring NMR spectra of metabolite samples by semiautomated process Basic Protocol 3: Measuring sample pH Support Protocol 1: Cleaning NMR tubes Basic Protocol 4: Processing raw spectra data Basic Protocol 5: Profiling spectra Support Protocol 2: Spectral profiling of sugars and other complex metabolites.

Adenosine to-inosine (A-to-I) RNA editing is a conserved post-transcriptional modification that is critical for a variety of cellular processes. A-to-I editing is widespread in nearly all types of RNA, directly imparting significant global changes in cellular function and behavior. Dysfunctional RNA editing is also implicated in a number of diseases, and A-to-I editing activity is rapidly becoming an important biomarker for early detection of cancer, immune disorders, and neurodegeneration. While millions of sites have been identified, the biological function of the majority of these sites is unknown, and the regulatory mechanisms for controlling editing activity at individual sites is not well understood. Robust detection and mapping of A-to-I editing activity throughout the transcriptome is vital for understanding these properties and how editing affects cellular behavior. However, accurately identifying A-to-I editing sites is challenging because of inherent sampling errors present in RNA-seq. We recently developed Endonuclease V immunoprecipitation enrichment sequencing (EndoVIPER-seq) to directly address this challenge by enrichment of A-to-I edited RNAs prior to sequencing. This protocol outlines how to process cellular RNA, enrich for A-to-I edited transcripts with EndoVIPER pulldown, and prepare libraries suitable for generating RNA-seq data. © 2020 Wiley Periodicals LLC.

Basic Protocol 1: mRNA fragmentation and glyoxalation

Basic Protocol 2: EndoVIPER pulldown

Basic Protocol 3: RNA-seq library preparation and data analysis