This month’s issue of CSH Protocols features an article by Andrew Salinger and Monica Justice, detailing a technique for Mouse Mutagenesis Using N-Ethyl-N-Nitrosourea (ENU) (article is freely available as one of our featured protocols). Back in the ancient days of my graduate school work, the idea of doing large scale forward genetics in mouse was unthinkable. Who had the space, let alone the funding and personnel to keep and track all of those cages? It was always one of those reasons we grumbled about the Drosophila labs, and the incredibly cool tools they had at their disposal. Over the years, the techniques were refined, and now, according to Justice, screens like this are an “established as part of a mouse geneticist’s toolkit,” and can be effective even in labs with very limited amounts of mouse space. So it’s nice to see this incredibly productive method readily available for use in mouse. Now if we can just do something about that pesky internal development that’s so limiting to imaging experiments…..

The March issue of CSH Protocols includes a diverse trio of methods for fluorescently labeling cells and subcellular structures. The most basic of the three methods comes from Brad Chazotte at Campbell University and covers labeling of lysozymes with Neutral Red. This joins a group of already-published protocols from Chazotte on labeling cellular structures including the plasma membrane, the golgi apparatus and acetylcholine receptors. Expect more articles in this series in forthcoming issues.

The second protocol provides a method for differentiating viable plant cells from dead plant cells. Contributed by Birgit Schwab and Martin Hülskamp from the Center for Plant Molecular Biology in Tübingen, the technique takes advantage of the inability of propidium iodide to enter live cells. Dead cells allow it in, and fluoresce red, so they can be easily identified.

Finally, Paul Kulesa’s group at the Stowers Institute have written up their method for Photoactivation Cell Labeling for Cell Tracing in Avian Development. This technique allows for selective marking of individual cells or groups of cells at precise times and spatial locations normally not accessible using previous techniques. It’s less invasive than most methods used for labeling cells in avian embryos, and can be targeted to both individual cells, or small groups of cells. This month’s cover image shows an example of this technique.

Immunohistochemistry (the localization of proteins in a tissue by binding antibodies to specific antigens) is a technique where one protocol definitely does not fit all. Each model organism seems to have its own quirks, whether it be in the fixative used, the methods needed for antibody penetration, issues with autofluorescence or even just figuring out which cross-species antibodies work in a given system. To that end, we’ve been working on expanding our coverage of immunohistological protocols. The February issue of CSH Protocols brings methods for plant sections, using both avidin-biotin and alk-phos, as well as a method for whole-mount immunocytochemistry in Xenopus embryos from John Wallingford’s lab at the University of Texas (they provided the lovely cover image for this month).

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Those familiar with the CSHL Press Manual, Drosophila Protocols (edited by Sullivan, Ashburner and Hawley) will want to be sure to check CSH Protocols’ February issue, as Bruce Paterson’s group at the National Institutes of Health has written an update of his chapter, “Targeted Disruption of Gene Function in Drosophila by RNA Interference”. The new, up-to-date version of the book chapter appears online in a series of articles, including Preparation of Double-Stranded RNA for Drosophila RNA Interference (RNAi), Collection of Drosophila Embryos for RNA Interference (RNAi), and Injection of dsRNA into Drosophila Embryos for RNA Interference (RNAi) (freely available as one of this month’s featured protocols). Paterson’s group has also contributed a new article covering Drosophila RNA Interference (RNAi) Using a Gal-4 Inducible Transgene Vector.

January’s issue of CSH Protocols is now available online, and it contains a set of protocols from Cathy Krull’s lab at the University of Michigan. The articles provide methods for electroporating your gene of interest into somites, neural crest cells and motor neurons. The accessibility of the chick embryo has long made it a standard model organism for developmental biology, and methods like these greatly enhance our abilities to tag and track cells, as well as to genetically manipulate the embryo. They’re even valuable for labs not working with avian systems, particularly mouse labs, because they offer the opportunity to get a quick and easy look at expression and potential effects of experimental constructs. Unlike making a transgenic mouse, an expensive and time-consuming process, working with chick eggs is inexpensive, and relatively rapid. Testing your mouse constructs in the chick embryo is a great way to fine tune the constructs themselves to ensure proper expression. It can also give insight into potential effects of construct expression, which can save valuable time once your transgenic mice are available, as you may already know where to start analyzing.

Fate-mapping, the tagging of specific cells or tissues in an embryo, and following their movements and development over time, has a long history as a valuable method. The earliest fate-maps date back to the 1880’s. The first “modern” fate-maps were created in 1929 by Walter Vogt, who applied vital dyes to regions of the amphibian embryo. This allowed him to track which embryonic regions developed into which adult tissues. Two methods, featured in the December issue of CSH Protocols and freely available to non-subscribers, present new fate-mapping techniques, which overcome some serious experimental barriers.
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In situ hybridization of mRNA has long been a standard laboratory practice. Over recent years, the technique has evolved from the laborious sectioning of tissues and treatment with radioactive probes to the easier colorimetric (and occasionally fluorescent) methods now in use. Recent issues of CSH Protocols featured articles detailing in situ hybridization in Xenopus, Drosophila (here as well), and cultured cells.

November’s issue of CSH Protocols provides a full set of instructions for in situ hybridization on mouse embryos, as well as a cutting-edge method for imaging real-time gene expression in living systems with single-transcript resolution.

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I know I’m a week or so late, but congratulations are in order for Martin Evans, Oliver Smithies and Mario Capecchi for winning the 2007 Nobel Prize in Physiology or Medicine “for their discoveries of principles for introducing specific gene modifications in mice by the use of embryonic stem cells”. Read the rest of this entry »

This month’s issue includes a protocol for building incubators for use in observing development. Read the rest of this entry »

One of June’s featured freely available protocols on CSH Protocols details the use of Keller explants and sandwiches, a tremendously useful Xenopus method originally developed to allow observation of gastrulation movements, particularly convergent extension. The key is that these explants remain flat instead of curling up, and the mesoderm elongates in a plane with the adjacent ectoderm, rather than involuting. This allows you to see what would normally be happening beneath the surface in an opaque embryo. Read the rest of this entry »