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.

While it is possible to analyze the global lipid composition of a cell, a deeper understanding of what lipids are doing within that cell is more difficult to come by. Though the lipid components may be known, finding their exact position, how dynamically they change location, and how rapidly they are metabolized presents an experimental challenge. The obvious approach would be the addition a fluorescent tag, which would allow for imaging of lipids in cells. Unfortunately, most commonly used fluorescent tags are as large as the lipid itself and are likely to have a strong effect on lipid location and metabolism.

In the July issue of Cold Spring Harbor Protocols, Joachim Goedhart and colleagues present a suite of protocols to get around these problems and allow for live imaging of lipids in cells. Their introduction to the topic explains the approach:

To circumvent this problem, two solutions have been developed–namely, the use of fluorescently labeled proteins that specifically recognize lipids and a chemical method to introduce the fluorescent tag inside the cell.

Protocols are provided for Transfection of Cells with DNA Encoding a Visible Fluorescent Protein-Tagged Lipid-Binding Domain, Labeling Lipids for Imaging in Fixed Cells, and Labeling Lipids for Imaging in Live Cells.

The zebrafish (Danio rerio) has rapidly become a favored model organism for studying developmental biology. One of the most commonly used methods for genetic manipulation in the zebrafish is the delivery of plasmids or oligonucleotides to cells within the living embryo via electroporation. When cells are exposed to brief electrical fields, transient membrane destabilization occurs and nucleic acids can cross the plasma membrane. When the electrical field is removed, the membrane seals and the nucleic acids are trapped inside the cell. In vivo electroporation has proven particularly effective for delivering fluorescent protein expression vectors for imaging and loss-of-function reagents such as morpholinos or RNA interference (RNAi) constructs for the knockdown of gene function. In the July issue of Cold Spring Harbor Protocols, Jack Horne and colleagues present Targeting the Zebrafish Optic Tectum Using In Vivo Electroporation, a modification of the technique that can be used to specifically target the developing optic tectum, the midbrain’s visual processing center. Instructions are given for the construction of electroporation electrodes, preparation and injection of DNA, and electroporation of the DNA into the embryonic brain.

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.

The June issue of Cold Spring Harbor Protocols includes an early preview of CSHL Press’ forthcoming RNA: A Laboratory Manual. Protocols covering basic RNA techniques are now available, including methods for purification of RNA by by SDS Solubilization and Phenol Extraction and by Using TRIzol (TRI Reagent), Ethanol Precipitation of RNA and the Use of Carriers, Preparation of Cytoplasmic and Nuclear RNA from Tissue Culture Cells, Removal of Ribosomal Subunits (and rRNA) from Cytoplasmic Extracts before Solubilization with SDS and Deproteinization, Removal of DNA from RNA, Nondenaturing Agarose Gel Electrophoresis of RNA and Polyacrylamide Gel Electrophoresis of RNA.

The last two on that list cover gel electrophoresis, two of the most important and frequently used techniques in RNA analysis. Electrophoresis is used for RNA detection, quantification, purification by size and quality assessment. Gels are involved in a wide variety of methods including northern blotting, primer extension, footprinting and analyzing processing reactions. The two most common types of gels are polyacrylamide and agarose. Polyacrylamide gels are used in most applications and are appropriate for RNAs smaller than approximately 600 nucleotides (agarose gels are preferred for larger RNAs). Polyacrylamide Gel Electrophoresis of RNA describes how to prepare, load and run polyacrylamide gels for RNA analysis. The is featured in the June issue, and as one of our featured articles, the full-text version is available to subscribers and non-subscribers alike.

This set is just a small sampling of the manual’s contents, basic techniques from an early chapter. The full table of contents can be seen here.

The dynamic nature of biological processes has long been difficult to document, as researchers have been limited to static studies based on fixed specimens. Methods like immunocytochemistry or in situ hybridization can only provide accurate information on one organism at one particular time point. As Scott Fraser has remarked, it’s akin to trying to figure out the rules of football from looking at a set of still photographs taken during a game. But recent developments in imaging techniques, particularly the use of Green Fluorescent Protein (GFP) and its variants, have provided nondestructive ways to study dynamic processes over time, taking our understanding into the fourth dimension.

These new imaging techniques generate an enormous amount of digital image data, which can be difficult to cope with as it builds up over time. Computer-based image analysis is required for the extraction of reproducible and quantitative information. Previously, Cold Spring Harbor Protocols has featured Khuloud Jaqaman and Gaudenz Danuser’s case study using particle tracking to study cellular dynamics. In the June issue of the journal, Roland Eils and colleagues present Tracking and Quantitative Analysis of Dynamic Movements of Cells and Particles. The article sketches a general workflow for quantitative analysis of live cell images and details automated methods for image analysis including preprocessing, segmentation, registration, tracking and classification.

The generation of transgenic plants can be a lengthy and difficult process. Transient expression assays have been developed as faster and more convenient alternatives for investigating gene function. These assays often take advantage of the ability of Agrobacterium to transfer foreign DNA into plant cells with intact cell walls. Agrobacterium-mediated transformation is, however, inefficient and shows great variability. In the May issue of Cold Spring Harbor Protocols, Andreas Nebenführ and colleagues from the University of Tennessee present FAST Technique for Agrobacterium-Mediated Transient Gene Expression in Seedlings of Arabidopsis and Other Plant Species, a quick, efficient and economical assay for gene function in intact plants. The technique involves cocultivation of young plant seedlings and Agrobacterium in the presence of Silwet-77. The Silwet-77 facilitates transformation, thus replacing a wounding or device-dependent vacuum step. As one of May’s featured articles, it is freely available to subscribers and non-subscribers alike.

Using a promoter that can drive expression at an appropriate level is crucial in designing constructs for gene expression. Promoters can be tested via transient or stable transfection. But transfection efficiency in such assays can be low, so promoters are commonly fused to heterologous reporter genes that encode enzymes that can be quantified using highly sensitive assays. The reporter protein’s activity or fluorescence within a transfected cell population is approximately proportional to the steady-state mRNA level. The May issue of Cold Spring Harbor Protocols includes updated versions of three commonly used assays for promoter strength.

The Luciferase Assay uses a gene from the firefly Photinus pyralis. This gene encodes a 61-kDa enzyme that oxidizes D-luciferin in the presence of ATP, oxygen, and Mg++, yielding a fluorescent product that can be quantified by measuring the released light with a luminometer. The luciferase assay is extremely rapid, simple, relatively inexpensive, sensitive, and possesses a broad linear range.

The Chloramphenicol Acetyltransferase Assay utilizes an Escherichia coli chloramphenicol acetyltransferase (CAT) reporter gene. CAT catalyzes the acetylation of [14C]chloramphenicol which is monitored by autoradiography following thin-layer chromatography (TLC). The percent conversion of [14C]chloramphenicol to acetyl-[14C]chloramphenicol can be measured by PhosphorImager analysis of the TLC plate, counting in a scintillation counter, or by densitometry analysis of an autoradiograph.

The Beta-Galactosidase Assay uses the E. coli lacZ gene which encodes a beta-galactosidase. Beta-gal activity is measured through a simple and inexpensive colorimetric assay. Cells are lysed and extracts are mixed with O-nitrophenyl-beta-D-galactopyranoside (ONPG), which results in a yellow product. The optical densities of the samples are then determined spectrophotometrically.

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 other blog where I write, The Scholarly Kitchen, has been nominated for a Webby, a fairly prestigious award in the online world. Since the winner is determined by the voting public, and since we’re up against some seriously stiff competition (including the NY Times and Wall Street Journal), your help would be greatly appreciated. You can vote for us here. Voting does require you to complete a short registration process. You can only vote once per email address. Voting ends April 29th, and results will be announced May 4th (a visual tutorial on how to vote can be found here).
From the Society for Scholarly Publishing:

On behalf of the SSP leadership, we are very proud that our community has generated a vehicle as widely recognized and effective as the Scholarly Kitchen. Now, we have an opportunity to increase the awareness of the SSP, the Scholarly Kitchen, and scholarly publishing in general.

Thanks in advance for your help!