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.

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.

Neurons are organized into anatomical and functional groups called “circuits”. The activity of these circuits is traditionally monitored using conventional electrophysiological techniques. But some cells, such as the submandibular ganglia, are difficult to impale for intracellular recordings. Instead, viral vectors can be used to deliver fluorescent calcium sensors for detecting activity in a living animal. Calcium Imaging of Neuronal Circuits In Vivo Using a Circuit-Tracing Pseudorabies Virus, from Lynn Enquist and colleagues at Princeton University, provides detailed instructions for the use of the pseudorabies virus (PRV) as a vector for imaging connectivity and activity of neuronal circuits. PRV has a broad host range but does not infect higher-order primates, and it travels along chains of synaptically connected neurons. The PRV strain used in this procedure encodes G-CaMP2, a sensitive fluorescent calcium sensor protein. Available in the April issue of Cold Spring Harbor Protocols, the method allows for reliable detection of endogenous circuit activity at single-cell resolution. As one of April’s featured articles, it is freely available to subscribers and nonsubscribers alike.

mRNA in situ hybridization is a standard laboratory technique for analyzing gene expression. In a small, transparent specimen like a zebrafish embryo, this technique is straightforward and works well. Cold Spring Harbor Protocols has a set of protocols (here, here and here) describing the method from Cecilia Moens. But what happens when you’re dealing with a larger, opaque zebrafish tissue like the adult brain? Unlike mammals, zebrafish exhibit intense ongoing neurogenesis in all areas of the central nervous system. Adult zebrafish are increasingly being used in behavioral studies as well. Because the number of antibodies useful for examining expression in zebrafish is limited, mRNA in situ hybridization is a vital tool for understanding what’s happening during these processes. In the February issue of Cold Spring Harbor Protocols, Reinhard Köster and colleagues from the Helmholtz Zentrum München provide an adaptation of the standard in situ method that deals with these larger, opaque tissues by staining them after vibratome sectioning, Analysis of Gene Expression by In Situ Hybridization on Adult Zebrafish Brain Sections. While the brain is used as the sample tissue in this protocol, it can easily be modified for analysis of other adult tissues.

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

As new imaging methods are developed and our knowledge of neural development deepens, methods for growing neuronal cells in culture where they are readily manipulated and observed become more and more important. In the August issue of Cold Spring Harbor Protocols, Amy MacDermott and colleagues from Columbia University provide a series of protocols describing neuronal culture techniques that are particularly useful for studying synapse formation and interconnection.

Preparation of Coverslips for Neuronal Cultures describes setting up glass coverslips for three different types of cell culture: Mass Culture, which results in an extensive and widespread network of synapses, Microisland Culture, which allows easier identification of synaptically connected neurons, and Macroisland Culture, which strikes a balance between the better survival rates and better identification of synapses offered by the two methods above.

Survival of CNS neurons in culture is usually greatly improved if the neurons are plated on top of a confluent astrocyte layer, as is described in Dissection, Plating, and Maintenance of Cortical Astrocyte Cultures. Under these conditions, neurons attach more firmly and develop a more easily identifiable bi-dimensional neuritic tree than in culture conditions where primary cells are plated on top of collagen, laminin or other cell-free substrates.

Once established the astrocyte layer can be used to in the Dissection, Plating, and Maintenance of Dorsal Horn Neuron Cultures, which are suitable for electrophysiological, molecular and immunocytochemical studies. Dorsal horn neurons can be grown by themselves, or co-cultured with embryonic dorsal root ganglion neurons as described in Dissection, Plating, and Maintenance of Dorsal Root Ganglion Neurons for Monoculture and for Coculture with Dorsal Horn Neurons.

May’s issue of Cold Spring Harbor Protocols saw the publication of the final two species from Volume I of our “Emerging Model Organisms” series. These articles have been collected and made available as a laboratory manual. June’s issue brings us the first species from Volume II, the Honeybee (Apis mellifera).

Because of their obviously important role in pollination, a great deal of recent research has gone into investigating diseases which affect honeybees, such as Colony Collapse Disorder. Bees also exhibit remarkable social behavior, complex learning and memory and language skills, making them an excellent system for neuroscience research into these topics. The haplo-diploid sex determination system of bees is also of great interest. The sequenced genome of honeybees has allowed for comparisons with other species, with some surprising results. The genes underlying circadian rhythms in bees are much more like those in mouse than those found in Drosophila. The same goes for DNA methylation in gene regulation, where bees, like mammals but unlike Drosophila, methylate DNA on CpG residues.

Protocols are provided for Fixation and Storage of Honeybee Tissues, Whole-Mount In Situ Hybridization of Honeybee Tissues, In Situ Hybridization of Sectioned Honeybee Tissues, Immunohistochemistry on Honeybee Embryos, and RNA Interference (RNAi) in Honeybee Embryos.

Each month’s issue of Cold Spring Harbor Protocols will feature new (and newly revisited) model organisms, and the next set will be collected in Volume II of the manual series, out some time early next year.

April’s issue of Cold Spring Harbor Protocols introduces, well, re-introduces two longstanding experimental model systems, the snail (Ilyanassa obsoleta) and the leech (Helobdella).

The use of Ilyanassa in the laboratory dates back to the 1890’s, and it was a favorite of Thomas Hunt Morgan’s in the 1930’s. As a member of the Lophotrochozoa, a group made up of nearly one third of the animal phyla, the snail exhibits a spiralian developmental program. In addition to its use to study spiral cleavage, Ilyanassa is also used to study asymetric cell division and other phenomena:

It is an important model for studies of metamorphosis, the ecology of parasitism, and imposex, a striking morphological disorder caused by the disruption of sexual endocrine systems by environmental contaminants. Ilyanassa is also useful for studies of comparative neurobiology.

Protocols are provided for Obtaining Embryos, Induction of Larval Metamorphosis, Fixation, Isolation of Genomic DNA, Protein Isolation, and Pressure Injection.

In the 1870’s, C.O. Whitman, director of the MBL at Woods Hole used a local leech species for developmental biology studies. Gunther Stent’s lab used leeches for neurobiology research in the 1970’s. Like Ilyanassa, leeches are Lophotrochozoans and exhibit spiral cleavage and thus are useful species for studying this poorly understood program of development. Leeches are also used for the study of segmentation, regeneration and neurogenesis.

Protocols are provided for Handling Embryos, Microinjection, Devitellinization, Silver Staining, Immunohistochemistry, In Situ Hybridization, and Preparation for Microscopy.

For more emerging (and re-emerging) model systems, these articles and others like them are collected in Volume 1 of a new laboratory manual series.