Single-Molecule Dynamics and Localization of DNA Repair Proteins in Cells.

Direct observation of individual protein molecules in their native environment, at nanometer resolution, in a living cell, in motion is not only fascinating but also uniquely informative. Several recent major technological advances in genomic engineering, protein and synthetic fluorophore development, and light microscopy have dramatically increased the accessibility of this approach. This chapter describes the procedures for modifying endogenous genomic loci to producing fluorescently tagged proteins, their high-resolution visualization, and analysis of their dynamics in mammalian cells, using DNA repair proteins BRCA2 and RAD51 as an example.

Genome organization in the nucleus: From dynamic measurements to a functional model.

A biological system is by definition a dynamic environment encompassing kinetic processes that occur at different length scales and time ranges. To explore this type of system, spatial information needs to be acquired at different time scales. This means overcoming significant hurdles, including the need for stable and precise labeling of the required probes and the use of state of the art optical methods. However, to interpret the acquired data, biophysical models that can account for these biological mechanisms need to be developed. The structure and function of a biological system are closely related to its dynamic properties, thus further emphasizing the importance of identifying the rules governing the dynamics that cannot be directly deduced from information on the structure itself. In eukaryotic cells, tens of thousands of genes are packed in the small volume of the nucleus. The genome itself is organized in chromosomes that occupy specific volumes referred to as chromosome territories. This organization is preserved throughout the cell cycle, even though there are no sub-compartments in the nucleus itself. This organization, which is still not fully understood, is crucial for a large number of cellular functions such as gene regulation, DNA breakage repair and error-free cell division. Various techniques are in use today, including imaging, live cell imaging and molecular methods such as chromosome conformation capture (3C) methods to better understand these mechanisms. Live cell imaging methods are becoming well established. These include methods such as Single Particle Tracking (SPT), Continuous Photobleaching (CP), Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Correlation Spectroscopy (FCS) that are currently used for studying proteins, RNA, DNA, gene loci and nuclear bodies. They provide crucial information on its mobility, reorganization, interactions and binding properties. Here we describe how these dynamic methods can be used to gather information on genome organization, its stabilization mechanisms and the proteins that take part in it.

Reliable measurement of the FRET sensitized-quenching transition factor for FRET quantification in living cells.

3-cube-based Förster resonance energy transfer (FRET) microscopy, a sensitized acceptor FRET quantification method, has been widely used to visualize dynamic protein-protein interaction in living cells. Determining the FRET sensitized-quenching transition factor (G factor) of a particular donor-acceptor pair and optical system is crucial for 3-cube FRET quantification. We here improved the acceptor photobleaching-based G factor determination method (termed as mPb-G) and the two-plasmid-based G factor determination method (termed as mTP-G) for rapid and reliable measurement of the G factor. mTP-G method determines G factor by simultaneously detecting three images of cells exclusively expressing each of two tandem constructs with multiple donors and multiple acceptors. This method circumvents switchover of the cells exclusively expressing each of the two constructs. mPb-G method images G factor by detecting three images of cells expressing a donor-acceptor tandem FRET construct before and after partially photobleaching acceptor. We performed the two methods on our dual-channel wide-field FRET microscope to obtain reliable G factor, and also measured the FRET efficiency and acceptor-to-donor concentration ratio of tandem constructs with different acceptor-donor stoichiometries in living HepG2 cells. mTP-G and mPb-G methods provide two simple and reliable tools for determining the G factor, in turn, quantitatively measuring FRET signal and monitoring dynamic biochemical processes in living cells.

Dissecting protein reaction dynamics in living cells by fluorescence recovery after photobleaching.

Proteins within most macromolecular complexes or organelles continuously turn over. This turnover results from association and dissociation reactions that are mediated by each of the protein's functional domains. Thus, studying organelle or macromolecular formation from the bottom up using theoretical and computational modeling approaches will necessitate the determination of all of these reaction rates in vivo. Yet current methods for examining protein dynamics either necessitate highly specialized equipment or limit themselves to basic measurements. In this protocol, we describe a broadly applicable method based on fluorescence recovery after photobleaching (FRAP) for determining how many reaction processes participate in the turnover of any given protein of interest, for characterizing their apparent association and dissociation rates, and for determining their relative importance in the turnover of the overall protein population. Experiments were performed in melanoma M2 cells expressing mutant forms of ezrin that provide a link between the plasma membrane and the cortical actin cytoskeleton. We also describe a general strategy for the identification of the protein domains that mediate each of the identified turnover processes. Our protocol uses widely available laser-scanning confocal microscopes, open-source software, graphing software and common molecular biology techniques. The entire FRAP experiment preparation, data acquisition and analysis require 3-4 d.

FRAP and FRET methods to study nuclear receptors in living cells.

Quantitative imaging techniques of fluorescently-tagged proteins have been instrumental in the study of the behavior of nuclear receptors (NRs) and coregulators in living cells. Ligand-activated NRs exert their function in transcription regulation by binding to specific response elements in promotor and enhancer sequences of genes. Fluorescence recovery after photobleaching (FRAP) has proven to be a powerful tool to study the mobility of fluorescently-labeled molecules in living cells. Since binding to DNA leads to the immobilization of DNA-interacting proteins like NRs, FRAP is especially useful for determining DNA-binding kinetics of these proteins. The coordinated interaction of NRs with promoters/enhancers and subsequent transcription activation is not only regulated by ligand but also by interactions with sets of cofactors and, at least in the case of the androgen receptor (AR), by dimerization and interdomain interactions. In living cells, these interactions can be studied by fluorescence resonance energy transfer (FRET). Here we provide and discuss detailed protocols for FRAP and FRET procedures to study the behavior of nuclear receptors in living cells. On the basis of our studies of the AR, we provide protocols for two different FRAP methods (strip-FRAP and FLIP-FRAP) to quantitatively investigate DNA-interactions and for two different FRET approaches, ratio imaging, and acceptor photobleaching FRET to study AR domain interactions and interactions with cofactor motifs. Finally, we provide a protocol of a technique where FRAP and acceptor photobleaching FRET are combined to study the dynamics of interacting ARs.

Determination of plasma membrane fluidity with a fluorescent analogue of sphingomyelin by FRAP measurement using a standard confocal microscope.

Membrane perturbing effects have been described in neurodegenerative process like Alzheimer's disease and prion disorders. For example, non fibrillar amyloid-beta peptides (Abeta) implicated in Alzheimer's disease may exert its toxicity via membrane perturbation. Membrane organisation can be evaluated by its influence on lateral diffusion of lipids, which itself can be measured by FRAP (fluorescence recovery after photobleaching). We used this technique to study the effects of Abeta on membrane fluidity (Pillot et al., manuscript in preparation). We propose here a simple adaptation of FRAP using standard confocal laser scanning microscopy (CLSM). As a test experiment, we analysed the lateral diffusion of a fluorescent analogue of sphingomyelin and were able to demonstrate its increase upon cholesterol depletion induced by methyl-beta-cyclodextrin (cdx).