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.

The use of recombinant proteins, antibodies, small molecules, or nucleic acids as affinity reagents is a simple yet powerful strategy to study the protein/bait interactions that drive biological processes. Analysis via mass spectrometry rather than western blotting extends the identification of interactors, often allowing detection of thousands of proteins from complex mixtures. But this increased sensitivity can lead to problems distinguishing specific interactions from background noise. In the March issue of Cold Spring Harbor Protocols, Shao-En Ong from the Broad Institute of MIT and Harvard presents Unbiased Identification of Protein/Bait Interactions Using Biochemical Enrichment and Quantitative Proteomics. This method uses quantitative proteomics approaches to compare enrichment with the bait of interest against samples using control baits to allow sensitive detection and discrimination of specific protein/bait interactions. As one of March’s featured articles, it is freely available to subscribers and non-subscribers alike.

The enzyme-linked immunospot (ELISPOT) assay is considered by many to be the gold standard for monitoring cellular immune responses. The method is highly sensitive, quantitative, easy to use and amenable to high throughput screening. Until recently, the ELISPOT assay has been limited to the characterization of only one single effector molecule. Since the maintenance of both IFN-gamma and IL-2 by pathogen-specific T cells has been linked to a more favorable clinical outcome in human immunodeficiency virus (HIV) and Leishmania infections, an ELISPOT assay able to characterize both these effector molecules would be helpful for monitoring immune responses to certain infectious agents. Nicole Bernard and colleagues from the McGill University Health Centre present a protocol for Dual-Color ELISPOT Assay for the Simultaneous Detection of IL-2 and/or IFN-gamma Secreting T Cells in the January issue of Cold Spring Harbor Protocols. As interest in multifunctional T-cell monitoring in human diseases grows, this method is likely to be extensively used. The protocol is one of January’s featured articles, and is freely available to subscribers and non-subscribers alike.

Live cell imaging techniques are driving a revolution in biological research. Instead of viewing dead tissues and cells fixed at a particular stage of activity, scientists can now visualize dynamic changes as they happen, permitting a better understanding of complete processes. The revolution has been fueled by the implementation of genetically encoded fluorescent proteins, the subject of the 2008 Nobel Prize in Chemistry.

The diverse array of applications benefiting from fluorescent proteins ranges from markers targeted at organelles and protein fusions designed to monitor intracellular dynamics to reporters of transcriptional regulation and in vivo probes for whole-body imaging and detection of cancer. Fluorescent proteins have enabled the creation of highly specific biosensors to monitor a wide range of intracellular phenomena, including pH and metal-ion concentration, protein kinase activity, apoptosis, membrane voltage, cyclic nucleotide signaling, and tracing neuronal pathways. In the December issue of Cold Spring Harbor Protocols, David Piston and colleagues present Fluorescent Protein Tracking and Detection: Fluorescent Protein Structure and Color Variants, a comprehensive overview of the wide variety of fluorescent proteins that are currently available. The article features more than twenty movies of different fluorescent proteins in action and is a great primer for planning imaging experiments. As one of December’s featured articles, it is freely available to subscribers and non-subscribers alike.

In addition, the same authors have also contributed Fluorescent Protein Tracking and Detection: Applications Using Fluorescent Proteins in Living Cells. This article provides some general tips for the practical aspects of using and imaging enhanced green fluorescent protein (EGFP) and newer members of the color palette, as well as quantitative imaging of fluorescent proteins and imaging of several fluorescent proteins at the same time. Finally, an overview is provided for the different types of biosensors that have been derived from flourescent proteins.

Both articles are adapted from the spectacular new manual, Live Cell Imaging: A Laboratory Manual, Second Edition which is due out by month’s end.

CSH Protocols December Cover