Laboratory Organisms

The Scuttle Fly (Megaselia abdita) (© 2011, CSHL Press)

Drosophila melanogaster, one of the best-studied genetic model systems in developmental biology, has a new laboratory companion:  the scuttle fly (Megaselia abdita).  The scuttle fly is phylogenetically located between Drosophila and mosquito.  It is easy to culture in the lab, and the embryos are suitable for functional studies.  For these reasons, Urs Schmidt-Ott (University of Chicago) has helped to develop the scuttle fly experimental system to assist his laboratory’s goals of understanding differences between Drosophila and mosquito development, and to help uncover genetic mechanisms of evolutionary change in insects.

In this month’s issue of Cold Spring Harbor Protocols, Schmidt-Ott and two of his lab members, Ab. Matteen Rafiqi and Steffen Lemke, have published a series of articles on the scuttle fly. Their introductory article includes background information about the species, advice on scuttle fly sources and husbandry, uses of the scuttle fly in evo-devo studies and in forensic entomology, and information about related species. While many standard Drosophila methods can be applied to scuttle fly, Schmidt-Ott and colleagues also wrote step-by-step protocols on maintaining cultures and collecting eggs, preparing embryos for injection, fixing and devitellinizing embryos, and preparing cuticles from injected embryos that have been specifically optimized for this species.

The scuttle fly is the latest addition to our ever-growing collection of emerging model organisms.  Click here for a complete list.

The cover of the March 2011 issue of Cold Spring Harbor Protocols, out today, features several striking images of mouse and quail embryos.  The method used to produce the images, microscopic magnetic resonance imaging (μMRI), is a noninvasive imaging technique that permits the visualization of regions deep within embryos that are inaccessible using optical methods.  During μMRI, the specimens remain in near-physiological conditions, remaining anatomically unperturbed.  The method is ideal, therefore, for generating developmental atlases of these organisms.

Quail embryos in a "relaxed" posture used to construct a μMRI-based developmental atlas. (©2011, CSHL Press)

In an accompanying article, the authors, Seth Ruffins and Russell Jacobs (Caltech Biological Imaging Center), describe the preparation of specimens for μMRI and appropriate applications of μMRI for developmental biology, including the construction of atlases.  Using these methods, they have successfully generated digital anatomical atlases of both quail and mouse development (see the Caltech MRI Atlases).  These atlases, and others constructed using μMRI, will be useful references for developmental biologists, providing identifiable anatomical landmarks and standards for comparison.

New technologies and methods are spurring a renaissance in the study of organogenesis. Organogenesis, essentially the process through which a group of cells becomes a functioning organ, has important connections to biological processes at the cellular and developmental levels, and its study offers great potential for medical treatments through tissue engineering approaches. The January issue of Cold Spring Harbor Protocols features a method from Washington University’s Hila Barak and Scott Boyle for Organ Culture and Immunostaining of Mouse Embryonic Kidneys. The kidney is particularly interesting as it also serves as a model for branching morphogenesis. The protocol describes the isolation, culture and fluorescent immunostaining of mouse embryonic kidneys. As one of January’s featured articles, the protocol is freely available to subscribers and nonsubscribers alike.

The “Brainbow” strategy was originally developed in mice, as a system for labeling neurons in a variety of different colors, allowing one to follow multiple cells regardless of their proximity. Brainbow uses a construct that carries sequences for red, blue and green fluorescent proteins in tandem array, with two pairs of lox sites flanking the first two fluorescent proteins. Recombination occurs in the presence of the Cre recombinase, and one gets a variety of outcomes, resulting in the production of a red, blue or green label. When more than one copy of the Brainbow cassette exists within a cell, the primary tones can be mixed, providing more possible color combinations. This provides a powerful platform for studying neuronal morphology and cell movements. In the January issue of Cold Spring Harbor Protocols, Alex Schier and colleagues offer Multicolor Brainbow Imaging in Zebrafish. This protocol translates the system for use in zebrafish, which offer the advantages of easy visualization of transparent embryos and efficient generation of labeled subjects.

For those looking to add to their arsenal of laboratory techniques, Cold Spring Harbor Laboratory Press has just released a new series of Imaging manuals.

I had a hand in putting these books together, and I’m always pleased when we manage to publish books that I know I would have found incredibly helpful in my previous incarnation as a bench scientist. These two hit home as I was a postdoc in an imaging lab. While that was ten (10!!!) years ago, it’s almost shocking to realize that there weren’t any comprehensive lab manuals out there that really covered the whole of bio-imaging, from the basics of optics to the most current, bleeding-edge techniques. Consider that problem solved, courtesy of series editor Rafa Yuste.

The new series spins off from a previous set of publications. In 2000, CSHL Press published Imaging Neurons, based on a CSHL laboratory course. The book was a few years ahead of its time, and the methods had really caught on by the time the sequel, Imaging in Neuroscience and Development was released in 2005. Five years later, and there’s been far too many new applications developed to fit into one volume, hence the release of the new series.

Imaging: A Laboratory Manual is the flagship of the series. It offers all the basics: optics, confocal, multi-photon, lasers, cameras, staining cells, etc. The manual goes on from there though, through labeling and indicators to advanced techniques like photoactivation, light sheet imaging, array tomography, fast imaging, molecular imaging, superresolution imaging and every acronym you can think of (FRET, FLIM, FRAP, FIONA, PALM, STORM, BiFC, AFM, TIRFM to name a sampling). If you have a microscope in your lab or if you spend any time in your local imaging center, this is the book you need.

Imaging in Developmental Biology: A Laboratory Manual is the second book in the series, just released. We old-school developmental biologists used to have to look at fixed sectioned specimens taken from different time points, and try to piece together the big picture of what was really happening as an embryo developed. New techniques have revolutionized our understanding of dynamic processes, as they allow for real-time imaging, often over the entire course of an organism’s development. Like the preceding volume, the book starts with the basics, methods for visualizing development in laboratory standard model organisms (C. elegans, Drosophila, zebrafish, Xenopus, avians and mouse) and then step by step brings the reader to the cutting edge of imaging technology.

The supplemental movies from both books are freely available through Cold Spring Harbor Protocols. Look for a third volume in March, on Imaging in Neuroscience, which will offer an astounding 90-plus chapters for analyzing every aspect of the nervous system in detail.

Visualizing mammalian development presents an obvious problem: embryos must develop in utero. That makes them a lot more difficult to see under a microscope than a zebrafish or a frog that develops as a free-standing egg. Extensive work has been done to develop embryonic culture techniques for external development of mouse embryos, allowing imaging approaches to be applied. Early efforts by members of Scott Fraser’s lab (including myself) provided a protocol for growing d 6.5-9.5 mouse embryos on the microscope stage. The December issue of Cold Spring Harbor Protocols features Imaging Cell Movements in Egg-Cylinder Stage Mouse Embryos from Oxford University’s Shankar Srinivas. The article describes a method for isolating and culturing much earlier mouse embryos, as well as an approach for time-lapse imaging as those embryos develop. While cell movements can be followed using light microscopy alone, the increasing variety of transgenic fluorescent reporter mice makes studies of cell movement easier and more informative. As one of December’s featured articles, the protocol is freely available to subscribers and non-subscribers alike.

New imaging technologies have revolutionized the study of developmental biology. Where researchers once struggled to connect events at static timepoints, imaging tools now offer the ability to visualize the dynamic form and function of molecules, cells, tissues, and whole embryos throughout the entire developmental process. In order to observe development over time, it is necessary to grow the embryos of laboratory model organisms on the microscope stage, and keep them as healthy and in as natural a state as possible. Methods for culturing and imaging the embryos of model organisms are featured in the December issue of Cold Spring Harbor Protocols.

Caenorhabditis elegans has been a key organism for understanding cellular differentiation and development. The fate of every one of the worm’s somatic cells has been mapped out, and its short developmental time, transparent shell, and nonpigmented cells makes C. elegans an ideal subject for imaging studies. Timothy Walston from Truman State University and Jeff Hardin from the University of Wisconsin-Madison provide An Agar Mount for Observation of Caenorhabditis elegans Embryos, an easy way to prepare live C. elegans embryos for microscopic visualization. The method involves embedding the embryo in agar to hold it in place,providing a fixed orientation for consistent imaging. Embryos prepared this way are amenable to both light microscopy and confocal microscopy. As one of our featured articles, the protocol is freely available to subscribers and non-subscribers alike.

Blood feeding mosquitoes transmit many of the world’s deadliest diseases, which are resurgent in developing countries and pose threats for epidemic outbreaks in developed countries. Recent mosquito genome projects have stimulated interest in the potential for disease control through the genetic manipulation of vector insects. To accomplish this, vector insects must be established as laboratory model organisms, allowing for a better understanding of their biology, and in particular, the genes that regulate their development. Aedes aegypti is a vector mosquito of great medical importance because it is responsible for the transmission of dengue fever and yellow fever. In the October issue of Cold Spring Harbor Protocols, Molly Duman-Scheel and colleagues present an overview of the background, husbandry, and potential uses of Ae. aegypti as a model species. Protocols are provided for culturing and egg collection, fixation and tissue preparation, whole mount in situ hybridization, immunohistochemical analysis and RNA interference in Ae. aegypti. This methodology, much of which is applicable to other mosquito species, is useful to both the comparative development and vector research communities.

This article series marks the latest entrant in Cold Spring Harbor Protocols’ long-running series on Emerging Model Organisms.

A cell devotes a significant amount of effort to maintaining the stability of its genome, preventing the sorts of chromosomal rearrangements characteristic of many cancers. Assays that measure the rate of gross chromosomal rearrangements (GCRs) are needed in order to understand the individual genes and the different pathways that suppress genomic instability. In the September issue of Cold Spring Harbor Protocols, Richard Kolodner and colleagues from the University of California, San Diego’s Ludwig Institute for Cancer Research present Determination of Gross Chromosomal Rearrangement Rates, a genetic assay to quantitatively measure the rate at which GCRs occur in yeast cells. The assay measures the rate of simultaneous inactivation of two markers placed on a nonessential end of a yeast chromosome. This simple protocol for determining GCR mutation rates in a variety of genetic backgrounds coupled with a diversity of modified GCR assays has provided tremendous insight into the large numbers of pathways that suppress genomic instability in yeast and appear to be relevant to cancer suppression pathways in humans. As one of September’s featured articles, the full text protocol is freely available to subscribers and nonsubscribers alike.

The Drosophila neuromuscular junction (NMJ) provides a superb model system for investigating the cellular and molecular mechanisms of synaptic transmission. The NMJ is large, easily accessed and its genetics are well-characterized. It shares many structural and functional similarities to synapses in other animals, including humans. In the September issue of Cold Spring Harbor Protocols, Bing Zhang and Bryan Stewart present an essential set of primers for electrophysiological recording from the Drosophila NMJ. The issue contains a detailed explanation of the Equipment Setup necessary, as well as instructions for Fabrication of Microelectrodes, Suction Electrodes, and Focal Electrodes. Protocols for Electrophysiological Recording from a ‘Model’ Cell, Electrophysiological Recording from Drosophila Larval Body-Wall Muscles, Voltage-Clamp Analysis of Synaptic Transmission at the Drosophila Larval Neuromuscular Junction, and Focal Recording of Synaptic Currents from Single Boutons at the Drosophila Neuromuscular Junction are also included. These protocols are adapted from Drosophila Neurobiology: A Laboratory Manual. Based on Cold Spring Harbor Laboratory’s long-running course, this manual has rapidly become an important resource for any neuroscience lab.

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