Current protocols in chemical biology最新文献

O-GlcNAcylation is a posttranslational modification involving the addition of the single monosaccharide N-acetylglucosamine (GlcNAc) onto serine and threonine residues of intracellular proteins. Though O-GlcNAc is found on ∼1000 proteins in mammals, its specific function on individual substrates remains largely a mystery. To overcome this shortcoming, work has been put toward developing metabolic chemical reporters (MCRs) to label O-GlcNAcylated proteins for subsequent biochemical analysis. Typically, these MCRs are GlcNAc or GalNAc analogs functionalized with azide or alkyne handles. These unnatural sugar moieties can be metabolically incorporated directly on to protein substrates. The protocols outlined in this article describe how to use MCRs as tools for visualizing and identifying potentially O-GlcNAc modified proteins via in-gel fluorescence, Western blotting, and mass spectrometry. Taken together, MCR labeling provides a powerful tool to discover where and when substrates are O-GlcNAc modified. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Treatment of cells and CuAAC

Basic Protocol 2: In-gel fluorescence of labeled cell lysates (1 mg scale)

Basic Protocol 3: Enrichment of labeled proteins, trypsinolysis, and collection of peptides for proteomics

Basic Protocol 4: Proteomic identification of labeled proteins

Small-molecule drug discovery can be hindered by the formation of aggregates that act as non-selective inhibitors of drug targets. Such aggregates appear as false positives in high-throughput screening campaigns and can bedevil structure-activity relationships during compound optimization. Protocols are described for resonant waveguide grating (RWG) and dynamic light scattering (DLS) as microplate-based high-throughput approaches to identify compound aggregation. Resonant waveguide grating and dynamic light scattering give equivalent results for the compound test set, as assessed with Bland-Altman analysis. © 2019 The Authors.

Basic Protocol 1: Resonant waveguide grating (RWG) in 384-well or 1536-well plate format to detect compound aggregation

Basic Protocol 2: Dynamic light scattering (DLS) in 384-well plate format to detect compound aggregation

Over the past few decades, numerous examples have demonstrated that intrinsic disorder in proteins lies at the heart of many vital processes, including transcriptional regulation, stress response, cellular signaling, and most recently protein liquid-liquid phase separation. The so-called intrinsically disordered proteins (IDPs) involved in these processes have presented a challenge to the classic protein "structure-function paradigm," as their functions do not necessarily involve well-defined structures. Understanding the mechanisms of IDP function is likewise challenging because traditional structure determination methods often fail with such proteins or provide little information about the diverse array of structures that can be related to different functions of a single IDP. Single-molecule fluorescence methods can overcome this ensemble-average masking, allowing the resolution of subpopulations and dynamics and thus providing invaluable insights into IDPs and their function. In this protocol, we describe a ratiometric single-molecule Förster resonance energy transfer (smFRET) routine that permits the investigation of IDP conformational subpopulations and dynamics. We note that this is a basic protocol, and we provide brief information and references for more complex analysis schemes available for in-depth characterization. This protocol covers optical setup preparation and protein handling and provides insights into experimental design and outcomes, together with background information about theory and a brief discussion of troubleshooting. © 2020 by John Wiley & Sons, Inc. Basic Protocol: Ratiometric smFRET detection and analysis of IDPs Support Protocol 1: Fluorophore labeling of a protein through maleimide chemistry Support Protocol 2: Sample chamber preparation Support Protocol 3: Determination of direct excitation of acceptor by donor excitation and leakage of donor emission to acceptor emission channel.

Subtiligase is a powerful enzymatic tool for N-terminal modification of proteins and peptides. In a typical subtiligase-catalyzed N-terminal modification reaction, a peptide ester donor substrate is ligated onto the unblocked N terminus of a protein, resulting in the exchange of the ester bond in the donor substrate for an amide bond between the donor substrate and protein N terminus. Using this strategy, new chemical probes and payloads, such as fluorophores, affinity handles, cytotoxic drugs, and reactive functional groups, can be introduced site-specifically into proteins. While the efficiency of this reaction depends on the sequences to be ligated, a panel of mutants was recently developed that expands the scope of substrate sequences that are suitable for subtiligase modification. This article outlines the steps for applying subtiligase or specificity variants for both site-specific bioconjugation of purified proteins and for global modification of cellular N termini to enable their sequencing by tandem mass spectrometry. © 2020 by John Wiley & Sons, Inc.

Basic Protocol 1: Subtiligase-catalyzed site-specific protein bioconjugation

Support Protocol 1: Expression and purification of subtiligase-His₆

Support Protocol 2: Subtiligase substrate synthesis

Basic Protocol 2: Subtiligase N terminomics using a cocktail of subtiligase specificity mutants

Cover: In DeMeester et al. (https://doi.org/10.1002/cpch.74), PG biosynthesis begins with formation of UDP-NAM through MurA/B and UDP-NAG. Recycling enzymes AmgK/MurU provide another route to synthesize UDP-NAM with NAM as the building block. UDP-NAM is converted into Park's nucleotide through enzymes MurC-F. MraY links Park's nucleotide to the cell membrane, where MurG then glycosylates this Lipid I fragment to form Lipid II. MurJ transports Lipid II into the periplasmic space, where transglycosylases (TGase) and transpeptidases (TPase) further crosslink these molecules to form mature PG. NAM probes (blue) with bioorthogonal functionality at the 2-N position (X) or 3-lactic acid position (Y) are metabolically incorporated into PG through both recycling and biosynthetic machineries.

Small open reading frames (smORFs) encode previously unannotated polypeptides or short proteins that regulate translation in cis (eukaryotes) and/or are independently functional (prokaryotes and eukaryotes). Ongoing efforts for complete annotation and functional characterization of smORF-encoded proteins have yielded novel regulators and therapeutic targets. However, because they are excluded from protein databases, initiate at non-AUG start codons, and produce few unique tryptic peptides, unannotated small proteins cannot be detected with standard proteomic methods. Here,, we outline a procedure for mass spectrometry-based detection of translated smORFs in cultured human cells from protein extraction, digestion, and LC-MS/MS, to database preparation and data analysis. Following proteomic detection, translation from a unique smORF may be validated via siRNA-based silencing or overexpression and epitope tagging. This is necessary to unambiguously assign a peptide to a smORF within a specific transcript isoform or genomic locus. Provided that sufficient starting material is available, this workflow can be applied to any cell type/organism and adjusted to study specific (patho)physiological contexts including, but not limited to, development, stress, and disease. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Protein extraction, size selection, and trypsin digestion

Alternate Protocol 1: In-solution C8 column size selection

Support Protocol 1: Chloroform/methanol precipitation

Support Protocol 2: Reduction, alkylation, and in-solution protease digestion

Support Protocol 3: Peptide de-salting

Basic Protocol 2: Two-dimensional LC-MS/MS with ERLIC fractionation

Basic Protocol 3: Transcriptomic database construction

Alternate Protocol 2: Transcriptomics database generation with gffread

Basic Protocol 4: Non-annotated peptide identification from LC-MS/MS data

Basic Protocol 5: Validation using isotopically labeled synthetic peptide standards and siRNA

Basic Protocol 6: Transcript validation using transient overexpression

The immunoproteasome (iCP), a specific isoform of the proteasome's catalytic particle, is becoming an important protein complex of interest in various diseases. However, there is still much left to be learned about its activity in cells and how this can be altered by various endogenous conditions or with treatment with small molecules. Current strategies to investigate the iCP lack in their ability to be used in live, intact cells, limiting them to use in endpoint experiments. The iCP-selective probe presented here has been shown to be compatible with various live-cell assays, including monitoring iCP activity kinetically in a plate reader–based assay and observing single cells with confocal microscopy. A well-studied iCP-selective inhibitor, ONX-0914, has also been demonstrated to decrease the fluorescence signal of the iCP probe in both of these assays, showing its potential function in investigating small-molecule modulators of the iCP. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Synthesis of an immunoproteasome-selective peptide-peptoid hybrid probe

Basic Protocol 2: Expression of the immunoproteasome in A549 cells

Basic Protocol 3: Using the immunoproteasome probe to monitor activity in live cells with a fluorescence plate reader

Basic Protocol 4: Using the immunoproteasome probe to monitor activity in live cells with confocal microscopy

Bacterial cells utilize small carbohydrate building blocks to construct peptidoglycan (PG), a highly conserved mesh-like polymer that serves as a protective coat for the cell. PG production has long been a target for antibiotics, and its breakdown is a source for human immune recognition. A key component of bacterial PG, N-acetyl muramic acid (NAM), is a vital element in many synthetically derived immunostimulatory compounds. However, the exact molecular details of these structures and how they are generated remain unknown due to a lack of chemical probes surrounding the NAM core. A robust synthetic strategy to generate bioorthogonally tagged NAM carbohydrate units is implemented. These molecules serve as precursors for PG biosynthesis and recycling. Escherichia coli cells are metabolically engineered to incorporate the bioorthogonal NAM probes into their PG network. The probes are subsequently modified using copper-catalyzed azide-alkyne cycloaddition to install fluorophores directly into the bacterial PG, as confirmed by super-resolution microscopy and high-resolution mass spectrometry. Here, synthetic notes for key elements of this process to generate the sugar probes as well as streamlined user-friendly metabolic labeling strategies for both microbiology and immunological applications are described. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Synthesis of peracetylated 2-azido glucosamine

Basic Protocol 2: Synthesis of 2-azido and 2-alkyne NAM

Basic Protocol 3: Synthesis of 3-azido NAM methyl ester

Basic Protocol 4: Incorporation of NAM probes into bacterial peptidoglycan

Basic Protocol 5: Confirmation of bacterial cell wall remodeling by mass spectrometry

Identification and characterization of small molecule–protein interactions is critical to understanding the mechanism of action of bioactive small molecules. Photo-affinity labeling (PAL) enables the capture of noncovalent interactions for enrichment and unbiased analysis by mass spectrometry (MS). Quantitative proteomics of the enriched proteome reveals potential interactions, and MS characterization of binding sites provides validation and structural insight into the interactions. Here, we describe the identification of the protein targets and binding sites of a small molecule using small molecule interactome mapping by PAL (SIM-PAL). Cells are exposed to a diazirine-alkyne-functionalized small molecule, and binding interactions are covalently captured upon UV irradiation. An isotopically coded, acid-cleavable biotin azide handle is attached to the conjugated proteins using copper-catalyzed azide-alkyne cycloaddition. Biotin-labeled proteins are enriched for on-bead digestion and quantitative proteomics. Acid cleavage of the handle releases the bead-bound conjugated peptides for MS analysis and isotope-directed assignment of the binding site. © 2019 by John Wiley & Sons, Inc.

Basic Protocol 1: Generation of a small molecule–conjugated protein sample following treatment of live cells

Alternate Protocol: Generation of a small molecule–conjugated protein sample following treatment of cell lysate

Basic Protocol 2: Copper-catalyzed azide-alkyne cycloaddition functionalization and enrichment of labeled peptides

Support Protocol 1: Synthesis of acid-cleavable, isotopically coded biotin picolyl azide handle

Support Protocol 2: Monitoring enrichment by immunoblotting

Basic Protocol 3: Mass spectrometry analysis to identify interacting proteins and conjugation sites

Cover: In Papa and Shoulders (https://doi.org/10.1002/cpch.70), Schematic of DIRex recombineering. (A) In DIRex recombineering, intermediates containing the conditionally lethal gene (ccdB) are first selected for using kanamycin. The modification (Mod) is directly repeated in the homology arms. Thus, the direct and inverted repeats (IR) promote spontaneous excision to yield the final recombinant. Successful recombinants are selected for by removing arabinose. In the absence of arabinose, the ccdA antitoxin is no longer expressed, and ccdB then kills unmodified cells. (B) DIRex is hypothesized to promote spontaneous excision through hybridization between the two inverted repeats to form a hairpin during replication that brings the direct repeats into close proximity (Bzymek & Lovett, 2001; Näsvall, 2017). The direct repeats can then promote strand slippage during synthesis, which results in excision of one of the direct repeats and everything between the direct repeats. See e70.