While it always gives an editor a warm feeling to publish the newest groundbreaking protocol from the bleeding-edge of science, it often takes a good amount of time for these types of methods to trickle into mainstream use. Many labs don’t have access to the specialized equipment or reagents called for, and sometimes it takes a lot of thought to realize how a completely new assay can be applied to one’s own research. Keeping that in mind, it’s not surprising to see that on CSH Protocols, our methods that see the highest use from our readers are usually the most basic ones, the absolute classics common across all labs. As an example, Ethanol Precipitation of DNA is consistently on our list of most read articles every month.

Because readers are seeking out these basic methods, we’re putting special emphasis on getting more into our collection, and May’s issue has some good additions. Dany Adams contributes instructions for Making Solutions from Dry Chemicals and Hydrated Compounds, as well as Making and Diluting Stock Solutions. You can’t get any more basic than that. These articles come from Dany’s tremendously useful book, Lab Math and are great examples of why it’s a book you should hand to every new student as they enter the lab (along with At The Bench, our other primer for survival in the laboratory).

May also brings a nice set of histology protocols, again covering the basics, including Fixation and Permeabilization, Paraffin Embedding, Decalcifying, Preparing Slides and Coverslips and Sectioning. A protocol for Hematoxylin and Eosin Staining, one of the most common histological stains used is also available.

Expect more of these basic methods in future issues but don’t worry, we’re scouting along the bleeding edge as well.

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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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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Fluorescence Imaging with One Nanometer Accuracy (FIONA) is a method for localizing single fluorophores (or single groups of fluorophores) in the X-Y plane with high spatial resolution. In the October issue of CSH Protocols, Paul Selvin and colleagues provide a series of articles explaining the concepts behind FIONA (and several related techniques) as well as protocols for FIONA equipment set-up, chamber construction, imaging and data analysis. Read the rest of this entry »

October’s issue of CSH Protocols is now online. I’m very happy about one of the featured freely available protocols, Preparation of Rodent Hippocampal Slice Cultures. I’ve been digging through our Neuroscience Imaging protocols for months now, and have been unable to post many of them on CSH Protocols, because they deal with slice cultures. Each seems to start with an established culture and proceeds to explain an imaging technique that can be used with it. Each referenced the Stoppini et al., paper from 1991 that doesn’t seem to be available online, other than as an abstract. Luckily for us all, Michael Dailey and his lab at the University of Iowa were willing to write up a modern update of the method. They’ve included a very useful movie showing one of the tricker parts of the technique. With this in place, you can now expect more neuro-imaging to turn up in CSH Protocols over the next few months, including this month’s protocols for Imaging Microglia in Live Brain Slices and Slice Cultures, Imaging FM Dyes in Brain Slices and Imaging Zinc in Brain Slices.

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