Proteins and Proteomics

Unlike the genome, which is essentially identical in every somatic cell of an organism, the proteome varies across cells, and there is no self-replicating amplification mechanism for proteins like the polymerase chain reaction (PCR) for DNA. Because of this, methods that extract, separate, detect, and identify proteins from extremely small samples are needed.  In the current issue of Cold Spring Harbor Protocols, Duke University’s Erich Jarvis and colleagues provide such a method, Microproteomics: Quantitative Proteomic Profiling of Small Numbers of Laser-Captured Cells.  The protocol uses laser-capture microdissection to isolate pure cell populations from tissue sections and nanoscale liquid chromatography/tandem mass spectrometry to simultaneously identify and quantify hundreds of proteins from samples as small as 1000 cells. It is one of this month’s featured articles and is freely accessible here.

N-terminalomics is a high-throughput strategy for identifying proteins by selectively enriching for and sequencing their N-terminal peptides by mass spectrometry. In the November issue of Cold Spring Harbor Protocols, Samie Jaffrey and colleagues from Cornell University present a newly-developed N-terminalomic approach, N-CLAP (N-terminalomics by Chemical Labeling of the alpha-Amine of Proteins). N-CLAP: Global Profiling of N-Termini by Chemoselective Labeling of the alpha-Amine of Proteins describes the use of Edman chemistry to modify all of the amines in proteins, followed by the generation of a new unmodified amine at the N-terminus after the removal of the first amino acid by peptide bond cleavage. The alpha-amine at the protein N-terminus is labeled with a cleavable biotin affinity tag, which facilitates the downstream purification of the N-terminal peptides. Peptides are eluted by cleaving the biotin affinity tag and identified by tandem mass spectrometry (MS/MS). N-CLAP can be used for the identification of signaling peptides for mature proteins as well as for global profiling of cleavage events that occur during cell signaling, such as apoptosis.

Post-translational modifications of histones play an important role in regulating chromatin dynamics and function. One such modification, methylation, is involved in the regulation of the epigenetic program of a cell, determining chromatin structure, and regulating transcription. Methylation of histones occurs on both lysine and arginine residues, and until recently, was thought to be an irreversible process. The recent discovery of histone demethylases revealed that histone methylation is more dynamic than previously recognized. The October issue of Cold Spring Harbor Protocols features a set of methods from Keiichi Nakayama and colleagues from Kyushu University for investigating demethylase activity. The protocol, In Vitro Histone Demethylase Assay, describes two different in vitro histone demethylase enzyme reactions and three different methods for measuring histone demethylase activity. These methods can be applied to measuring histone demethylase activity in tissues and cell lysates, identification of novel histone demethylases, and screening for inhibitors of histone demethylases. As one of our featured articles, the protocol is freely available to subscribers and nonsubscribers alike.

Improvements in automation and acquisition time have made the microscope a viable platform for performing hundreds of concurrent parallel experiments. Using these sorts of tools, it is now possible to run high-throughput screens for protein function and interaction in living cells, examining dynamic cellular processes to distinguish between primary and secondary phenotypes, and to study the phenotype kinetics. In the August issue of Cold Spring Harbor Protocols, Jan Ellenberg and colleagues from the EMBL present High-Throughput Microscopy Using Live Mammalian Cells, an overview of how to screen live cells using imaging technologies. The article examines each aspect of the general screening process and considers specific examples in the processing of time-lapse experiments. The techniques discussed are based on the use of cultured mammalian cells, but the concepts are easily transferred to cultured cells from other species like Drosophila and small organisms such as C. elegans.

Immunoprecipitation is a commonly used technique for isolating and purifying a protein of interest. An antibody for the protein is incubated with a cell extract, and the resulting antibody/antigen complex is pulled out of solution. The method used for preparation of the cell extract can be critical for the experiment’s success. The choice of lysis conditions must be tailored to the nature of the epitope recognized by the immunoprecipitating antibody. Lysis of Cultured Cells for Immunoprecipitation, featured in the August issue of Cold Spring Harbor Protocols provides instructions for the lysis of cells grown as monolayer cultures and cells grown in suspension. The protocol offers a detailed comparison between different commonly used lysis buffers and protease inhibitor cocktails, as well as a guide to preparing a general protease inhibitor cocktail. As one of our featured articles, the protocol is freely available to subscribers and non-subscribers alike.

Producing recombinant proteins in bacterial hosts is a widely-used laboratory procedure. But generating a large yield of protein is often challenging. Getting enough raw material for experiments can be a time-consuming and frustrating process. In the August issue of Cold Spring Harbor Protocols, Jianjun Wang and colleagues present a method for Preparation of Very-High-Yield Recombinant Proteins using Novel High-Cell-Density Bacterial Expression Methods. By combining traditional IPTG induction with high-cell-density auto-induction, the method routinely produces 15-35 mg of pure protein from 50 mL bacterial cell cultures. Detailed protocols are given for preparation of a starting culture, double colony selection and optimization of expression conditions, which ensure plasmid stability resulting in a high yield of recombinant protein production.

Cold Spring Harbor Laboratory Press’ new Drosophila Neurobiology laboratory manual covers the three main approaches taught in the CSHL course: studying neural development, recording and imaging the nervous system, and studying behavior. The featured electrophysiology paper is part of the recording/imaging section, while the second featured article in the July issue of Cold Spring Harbor Protocols comes from a neural development chapter.

The larval Drosophila brain has been a valuable model for investigating the role of stem cells in development. These neural stem cells, called “neuroblasts,” have provided insight into the role of cell polarity in influencing cell fate. Identifying neuroblasts and their progeny requires a method capable of recognizing cell polarity and cell fate markers. Immunofluorescent Staining of Drosophila Larval Brain Tissue, provided by Cheng-Yu Lee and colleagues, describes procedures for the collection and processing of Drosophila larval brains for analysis of these markers. Neuroblasts are identified via immunolocalization, the use of labeled antibodies that specifically bind the marker proteins of interest. As one of our featured articles, it is freely available to subscribers and non-subscribers alike.

April’s issue of Cold Spring Harbor Protocols includes instructions for Rapid Coomassie Blue Staining of Protein Gels. This method is an adaptation of the conventional Coomassie staining protocol described in Staining Proteins in Gels with Coomassie Blue. Coomassie Brilliant Blue R250 (CBR-250) is the most commonly used dye for visualizing proteins because of its relatively high sensitivity. The modified method speeds up the destaining process for faster results with increased sensitivity and is compatible with mass-spectrometry-based methods for identifying proteins. Other methods for staining proteins can also be found in Cold Spring Harbor Protocols, including the Zinc/Imidazole Procedure for Visualization of Proteins in Gels by Negative Staining, and Staining Proteins in Gels with Silver Nitrate. Silver Nitrate’s sensitivity is in the low-nanogram range, which is 50-100 times more sensitive than classical Coomassie Blue staining, ~10 times better than colloidal Coomassie Blue staining, and at least twice as sensitive as the zinc/imidazole negative staining method.

The baculovirus expression vector system has been widely used to produce proteins originating from both prokaryotic and eukaryotic sources. It offers easy cloning techniques and abundant viral propagation, and since it is based on an insect cell environment, it provides eukaryotic posttranslational modification machinery. Surface modifications of the viral capsid enable specific targeting. Such modifications can be used to enhance viral binding and entry to a wide variety of both dividing and nondividing mammalian cells, as well as to produce antibodies against the displayed antigen. In addition, the technology should enable modifications of intracellular behavior, i.e., trafficking of recombinant “nanoparticles,” a highly relevant feature for studies of targeted gene or protein delivery. In the March issue of Cold Spring Harbor Protocols, Christian Oker-Blom and colleagues provide a suite of articles detailing the use of baculovirus-based display and gene delivery systems. Their protocol for Creation of Baculovirus Display Libraries is a featured article for March, and is freely available, along with nearly 90 other featured articles.

The recent explosion in the availability and variety of fluorescent proteins, new organic dyes and quantum dots has been a driving force in the growing use of Total Internal Reflection Fluorescence Microscopy (TIRFM). TIRFM only illuminates molecules that are within a thin volume near the coverslip surface of a specimen and not those deeper in solution. This allows for an unparalleled signal-to-noise ratio and tremendous resolution. In the March issue of Cold Spring Harbor Protocols, Samara Reck-Peterson, Nathan Derr and Nico Stuurman present Imaging Single Molecules Using Total Internal Reflection Fluorescence Microscopy (TIRFM), which includes an overview of the theory behind TIRFM, considerations for TIRFM setup and purification/labeling of proteins, and a discussion of new techniques for imaging single molecules with super-resolution localization. In addition, the group offers step-by-step protocols for Determining Single-Molecule Intensity as a Function of Power Density and Imaging Single Molecular Motor Motility with TIRFM. An example of TIRFM imaging of single dynein molecules labeled with TMR (green) moving along axonemal microtubules labeled with Cy5 (red) can be seen here.

Next Page »