Whole Embryo Chip

Chromatin immunoprecipitation would be a powerful tool for deciphering transcriptional regulation during development. However, most ChIP studies have involved cultured cells. On p. 34, Havis et al. describe a ChIP assay that can be used with whole zebrafish embryos. The method, which can be used up to 12 hours postfertilization, uses a pronase digestion for dechorionation, followed by homogenization and centrifugation. The pelleted nuclei are treated with formaldehyde for DNA-protein cross-linking, then lysed, and the chromatin is fragmented by sonication. The authors describe the appropriate conditions for adequate shearing of the chromatin and show that subsequent ChIP yields the expected results for antibodies against acetylated histone H4. Havis et al. further demonstrate the efficacy of their system by successful ChIP analysis of a transcription factor, in this case, exogenously introduced Myc-tagged Meis. Given the importance of Danio rerio in developmental biology and drug discovery, a well-optimized ChIP protocol for zebrafish embryos should have wide application in mapping target genes of transcription factors.

They Glow Together

Bimolecular fluorescent complementation (BIFC) describes a useful trait of certain recombinant fluorescent proteins that allows them to be expressed as two separate nonfluorescing units, which, when brought together, create a strong signal under laser light. This flexible system allows researchers to attach the two complementary halves of these molecules to separate proteins, providing a convenient and facile assay for detecting whether these proteins interact in cultured cells. Previous experiments of this type have been limited by diminished fluorescence at physiological temperatures, requiring lower temperature preparatory incubations that potentially negatively impacted the data collected. In this paper by Shyu et al. (p. 61), data is presented utilizing newly derived combinations of the N- and C-terminal domains from the recently characterized proteins Venus and Citrine—both mutants of enhanced yellow fluorescent protein (EYFP)—that function significantly better under true physiological conditions than their parent. The complementary halves of these new fluors were linked to the basic leucine zipper regions of c-Jun and c-Fos (b-Jun and b-Fos, respectively) and used in a variety of combinations to demonstrate that, in addition to being amenable to incubation at 37°C, they also provided a stronger signal (up to 15-fold) and required a shorter incubation time than the commonly used EYFP. Furthermore, improved multicolor BiFC could be achieved using an enhanced cyan fluorescent protein (ECFP) derivative, named Cerulean, when coupled with the complementary region from its parent or from Venus.

Green Screen

Almost as fast as scientists can come up with treatments, the disease-causing organisms that they are targeting seem to evolve mechanisms to resistant their eradication. This is true not only for bacteria, for which multidrug resistance is becoming an increasingly grave problem, but also for viruses such as HIV. Scientists and medical practitioners are in a constant battle with evolving resistance, obliged to come up with new and increasingly potent drugs to keep life-threatening diseases under control. Unfortunately, it is frequently true that the more powerful the drug, the more severe the side-effects. Technologies that allow more rapid and less expensive high-throughput screening are therefore both desirable and necessary. Published in this issue on p. 91, Ochsenbauer-Jambor et al. describe a newly-derived T cell line that expresses both HIV-1 co-receptors CXCR4 and CCR5 and also carries a stably integrated EGFP reporter gene that is up-regulated on infection with HIV-1. The former characteristic makes this cell line susceptible to infection by a wider range of HIV strains than other lines currently in use, while the latter makes it possible to screen clones more rapidly using fluorescent microscopy, without further manipulation of the cells or addition of other reagents. The improved dynamic range of the EGFP reporter—and the particular and unique characteristics of this cell line—make it exceptionally amenable to high-throughput assays using a 384-well format and thus a valuable tool for rapid screening of HIV-1 entry inhibitors and the development of HIV vaccines.

Whole Mount Brdu Staining

Assessing cellular proliferation in whole-mount preparations is typically done using BrdU labeling. Unfortunately, traditional methods usually rely upon treatment with hydrochloric acid in order to expose the BrdU-labeled epitopes to the antibody. This harsh procedure can significantly alter specimen morphology and denature protein epitopes that might otherwise be recognized by other antibodies in double-labeling experiments. At the same time, this treatment rarely allows efficient penetration of antibodies beyond the superficial cell layers in the specimen, resulting in inconsistent or misleading staining patterns. As an alternative, Tkatchenko (p. 29) explored whether DNase I treatment—commonly used in cells prepared for flow cytometric analysis and adapted to use in tissue sections—could be applied to BrdU labeling in whole mounts. Using retina quadrants as whole mounts, the author demonstrates enviable BrdU staining, including of cells deep within the specimen. At the same time, the samples display excellent morphology and are fully compatible with double labeling applications. With minimal optimization, the method should be equally useful for whole-mount embryos or thick vibratome slices of adult tissue.

Straightforward Tandem Epitope Tagging

Elucidating transcriptional regulatory networks in bacteria is an interesting and rewarding endeavor; generating the many requisite bacterial strains with different epitope-tagged proteins is not. Happily, Cho et al. (p. 67) have developed a painless strategy for tandem-epitope tagging based on PCR and two recombination systems. Multiple epitope tags are important for situations in which the tagged protein is present at low levels (transcription factors, for instance), while Red- and Flp/FRT-mediated recombination are widely used in recombineering and removing selectable markers, respectively. The authors' strategy rests upon a series of newly constructed template plasmids: each contains a multimeric tag, a stop codon, and an FRT-flanked kanamycin resistance marker. Users need simply PCR this cassette using gene-specific primers and then transform into a Red recombination proficient bacterial strain. Following homologous recombination, the cassette will be integrated into the chromosome; a subsequent site-specific recombination will result in excision of the selectable marker. In total, four template plasmids were created, containing two, four, six, or eight copies of the Myc epitope, respectively. As expected, the signal measured by Western blot analysis increased proportionally to the number of tags. The authors further confirmed that the multimeric tags did not significantly affect the function of tested target genes and demonstrated the utility of these tagged proteins for ChIP experiments. This ready-to-use, validated system should be of utility to researchers looking for a clean, convenient system for efficient protein tagging in E. coli.

Going with the Flow

To make bedside and field diagnostics a reality, rather than just a Hollywood movie subplot, requires the miniaturization of assay components that normally would take up a significant amount of any postdoc's desk space. Enter microfluidics. This particular field, at the intersection between physics, engineering, and biology, endeavors to provide a physical support that allows the organized and controlled mixing of assay components in nanoliter volumes, sufficiently small to engineer into a handheld device. In the report on p. 85 of this issue, Kartalov et al. describe a prototype 100-chamber microfluidic device capable of carrying out up to 5 tests on 10 samples. The polydimethylsiloxane chip was fabricated by soft lithography to contain a network of channels through which the reagents are pushed, their course controlled by a system of micromechanical valves. In each so-called capture microchamber, monoclonal antibodies to a single plasma protein are covalently attached to the chamber surface. Blood samples are then driven over the antibodies, allowing for the binding of the corresponding antigen. Following washing, biotinylated antibodies are passed over the resulting immunostack, the binding of which is detected by fluorescence of an Alexa Fluor®-conjugated streptavidin label. Assays could be carried out within a clinically relevant range using dramatically less sample volume (100 nL) and less antibody volume (300 nL) than current top-of-the-line systems, making this device a significant step forward in microfluidic research.