Large segments of DNA can vary in copy number between individuals. Such copy number variations (CNVs) contribute greatly to genetic diversity and are also thought to be associated with susceptibility or resistance to some diseases, including cancer. Simple Copy Number Determination with Reference Query Pyrosequencing (RQPS), featured in the September issue of Cold Spring Harbor Protocols, provides an assay for determining the copy number of any allele in the genome. The method, from Raphael Kopan and colleagues at Washington University, takes advantage of the fact that pyrosequencing can accurately measure the ratio of DNA fragments in a mixture that differ by a single nucleotide. A reference allele with a known copy number and a query allele with an unknown copy number are engineered with single nucleotide variations, and the ratio seen between these probes and genomic DNA reflects the copy number. RQPS can be used to measure copy number of any transgene, differentiate homozygotes from heterozygotes, detect the CNV of endogenous genes, and screen embryonic stem cells targeted with bacterial artificial chromosome (BAC) vectors. RQPS is rapid, inexpensive, sensitive, and adaptable to high-throughput approaches. As one of our featured articles, the protocol is freely available to subscribers and non-subscribers 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.

Zinc finger nucleases (ZFNs) are artificial restriction enzymes made by fusing an engineered zinc finger DNA-binding domain to the DNA cleavage domain of a restriction enzyme. ZFNs can be used to generate targeted genomic deletions of large segments of DNA in a wide variety of cell types and organisms. In the August issue of Cold Spring Harbor Protocols, Jin-Soo Kim and colleagues present Analysis of Targeted Chromosomal Deletions Induced by Zinc Finger Nucleases, a detailed protocol for the detection and analysis of large genomic deletions in cultured cells introduced by the expression of ZFNs. The method described allows researchers to detect and estimate the frequency of ZFN-induced genomic deletions by simple PCR-based methods. As one of our featured articles, the protocol is freely available to subscribers and non-subscribers alike.

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.

Since the early days of the 20th century and Thomas Hunt Morgan’s famous “Fly Room” at Columbia University, the fruit fly Drosophila melanogaster has been at the forefront of biological research. The powerful arsenal of experimental methods developed for this model organism is now being used to tackle one of the great scientific challenges of a new century: understanding the nervous system. Cold Spring Harbor Laboratory’s Neurobiology of Drosophila course has served as the training ground for a generation of scientists tackling these complex problems. A new laboratory manual based on the protocols and background information taught in the course promises to spread these techniques to a wider audience. Methods from the manual are featured in the July issue of Cold Spring Harbor Protocols.

When a fly is confronted with danger, it jumps into the air and flies away. The giant fiber system (GFS) of Drosophila is a neuronal circuit that mediates this escape response. The neurons in the GFS are readily identified and easily accessible for experimental assay. Electrophysiological Recordings from the Drosophila Giant Fiber System, from Marcus Allen and Tanja Godenschwege, describes a simple procedure for stimulating neurons directly in the brain of the adult fly and obtaining recordings from the output muscles of the GFS. As one of our featured articles, the protocol is freely available to subscribers and nonsubscribers 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.