Current Protocols in Protein Science最新文献

Tryptic peptide imaging is a primary workflow for matrix-assisted laser desorption/ionization imaging mass spectrometry (MALDI IMS) and has led to new information reporting highly multiplexed protein localization. Technological advances within the last few years have produced robust tools for automated spraying of both matrix and enzymes. When combined with high-mass-resolution and high-mass-accuracy instrumentation, studies now generally result in two-dimensional mapping of well over 1,000 peptide peaks. This protocol describes sample preparation, spraying, and application of enzymes and matrices, and MALDI FT-ICR instrumental considerations for two-dimensional mapping of tryptic peptides from fresh-frozen or formalin-fixed, paraffin-embedded tissue sections. Procedures for extraction of tryptic peptides from tissue sections for LC-MS/MS identification are also described. © 2018 by John Wiley & Sons, Inc.

Protein-carbohydrate interaction is essential for biological systems, and carbohydrate-binding proteins (CBPs) are important targets when designing antiviral and anticancer drugs. Due to the high cost and difficulty associated with experimental approaches, many computational methods have been developed as complementary approaches to predict CBPs or carbohydrate-binding sites. However, most of these computational methods are not publicly available. Here, we provide a comprehensive review of related studies and demonstrate our two recently developed bioinformatics methods. The method SPOT-CBP is a template-based method for detecting CBPs based on structure through structural homology search combined with a knowledge-based scoring function. This method can yield model complex structure in addition to accurate prediction of CBPs. Furthermore, it has been observed that similarly accurate predictions can be made using structures from homology modeling, which has significantly expanded its applicability. The other method, SPRINT-CBH, is a de novo approach that predicts binding residues directly from protein sequences by using sequence information and predicted structural properties. This approach does not need structurally similar templates and thus is not limited by the current database of known protein-carbohydrate complex structures. These two complementary methods are available at https://sparks-lab.org. © 2018 by John Wiley & Sons, Inc.

Sequence similarity searching has become an important part of the daily routine of molecular biologists, bioinformaticians and biophysicists. With the rapidly growing sequence databanks, this computational approach is commonly applied to determine functions and structures of unannotated sequences, to investigate relationships between sequences, and to construct phylogenetic trees. We introduce arguably the most popular BLAST-based family of the sequence similarity search tools. We explain basic concepts related to the sequence alignment and demonstrate how to search the current databanks using Web site versions of BLASTP, PSI-BLAST and BLASTN. We also describe the standalone BLAST+ tool. Moreover, this unit discusses the inputs, parameter settings and outputs of these tools. Lastly, we cover recent advances in the sequence similarity searching, focusing on the fast MMseqs2 method. © 2018 by John Wiley & Sons, Inc.

Heparan sulfate (HS) plays an important role in development and disease. It interacts with many growth factors, chemokines, and other ligands known to be important for cell growth, motility, and differentiation. However, isolating an antibody to HS in mice, rabbits, or humans is difficult due to the poor immunogenicity of HS. Phage display is a major antibody engineering technology that allows the selection of antibodies for poorly immunogenic or highly conserved antigens. This protocol contains detailed procedures for HS antigen preparation and isolation of a phage displayed human single-chain Fv (HS20) that binds HS on glypican-3 (GPC3), and analysis of the selected phage antibody. It is conceivable that the procedures described in this protocol may be applicable to the isolation of antibodies for a variety of HS molecules. © 2018 by John Wiley & Sons, Inc.

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to investigate subunit composition, track post-translational modifications, and verify identity and homogeneity of protein samples. It can also serve to purify proteins for use in further applications. In polyacrylamide gel electrophoresis, proteins migrate in response to an electrical field through pores in a polyacrylamide gel matrix; pore size decreases with increasing acrylamide concentration. Nondenaturing or “native” electrophoresis—i.e., electrophoresis in the absence of denaturants such as detergents and urea—is an often-overlooked technique for determining the native size, subunit structure, and optimal separation of a protein. Because mobility depends on the size, shape, and intrinsic charge of the protein, nondenaturing electrophoresis provides a set of separation parameters distinctly different from mainly size-dependent denaturing sodium dodecyl sulfate—polyacrylamide gel electrophoresis and charge-dependent isoelectric focusing. Two protocols are presented below. Continuous PAGE is highly flexible, permitting cationic and anionic electrophoresis over a full range of pH. The discontinuous procedure is limited to proteins negatively charged at neutral pH but provides high resolution for accurate size calibration.

Glycosylation of cell surface, secreted, and circulating proteins is one of the most common types of post-translational modification. These modifications occur most commonly as one of three major classes: N-linked glycosylation on asparagine residues, O-linked glycosylation on serine or threonine residues, or as glycosaminoglycan oligosaccharide polymers on serine. Specifically, for N-linked glycans, an endoglycosidase enzyme, peptide N-glycosidase F (PNGase F), cleaves the attached oligosaccharides between the asparagine and first sugar. A method to analyze released N-glycans and map them to specific locations within a tissue is presented here. The PNGase F is applied by solvent sprayer as a molecular layer on frozen or formalin-fixed tissues and all released N-glycans in a given region of tissue are detected using matrix-assisted laser desorption/ionization (MALDI) imaging mass spectrometry (MALDI-IMS). Using the described MALDI-IMS protocol, at least 40 or more individual N-glycans can be mapped to tissue histopathology and extracted for further structural analysis approaches. © 2018 by John Wiley & Sons, Inc.

Methods to efficiently deliver fluorophores across the cell membrane are crucial for imaging the dynamics of intracellular proteins using fluorescence. Here we describe a simple protocol for permeabilizing living cells using streptolysin O, a bacterial toxin, which allows transient uptake of fluorescent probes for labeling specific intracellular proteins. The technique is applicable for delivering different classes of fluorescent probes with a molecular weight of <150 kDa, and it is also applicable to a variety of different cell lines. The technique enables the utilization of a broad range of fluorophores for live cell imaging of intracellular proteins. Extended observation of intracellular fluorescence bound to specific proteins is now possible through super-resolution microscopy by using fluorophores that are photostable in “cell-friendly” deoxygenating and reducing conditions. © 2018 by John Wiley & Sons, Inc.

Phos-tag gels are recent tools to dissect protein phosphorylation that operate by inducing a shift in the electrophoretic mobility of phosphorylated proteins compared to their nonphosphorylated counterparts. This article describes the preparation and electrophoresis of Zn²⁺-Phos-tag gels along with electrotransfer of the separated phospho- and nonphosphoproteins onto a PVDF membrane using either wet-tank or semidry transfer. We also discuss the theory behind the technology with critical parameters to keep in mind for its successful application. © 2018 by John Wiley & Sons, Inc.

Membrane protein studies usually require use of detergents to extract and isolate proteins from membranes and manipulate them in a soluble context for their functional or structural characterization. However, solubilization with detergent may interfere with MP stability and may directly affect MP function or structure. Moreover, detergent properties can be affected such as critical micellar concentration (CMC) can be affected by the experimental conditions. Consequently, the experimenter must pay attention to both the protein and the behavior of the detergent. This article provides a convenient protocol for estimating the CMC of detergents in given experimental conditions. Then, it presents two protocols aimed at monitoring the function of a membrane protein in the presence of detergent. Such experiments may help to test various detergents for their inactivating or stabilizing effects on long incubation times, ranging from few hours to some days. © 2018 by John Wiley & Sons, Inc.

Determining ligand binding kinetics provides an indirect route to probe the functional capabilities of the binding pocket of a membrane protein receptor. Presented in this unit are four ligand-binding protocols that provide data useful for characterizing membrane proteins, including equilibrium binding, thermostability, competitive ligand binding, and kinetic ligand binding. These techniques use fluorescence anisotropy, which is safer, less costly, and simpler to execute than radioactive ligand binding. Each protocol may be used on its own or in combination with others to quantify a number of ligand binding constants. © 2018 by John Wiley & Sons, Inc.