GFP fluorescence: A few lesser-known nuggets that make it work.

Use of Green Fluorescent Protein (GFP) as a marker has revolutionized biological research in the last few decades. In this brief commentary, we reflect upon the success story of GFP and highlight a few lesser-known facets about GFP that add up to its usefulness.

Monitoring the Mitochondrial Dynamics in Mammalian Cells.

Mitochondria exist in a dynamic state inside mammalian cells. They undergo processes of fusion and fission to adjust their shape according to the different cell needs. Different proteins tightly regulate these dynamics: Opa-1 and Mitofusin-1 and Mitofusin-2 are the main profusion proteins, while Drp1 and its different receptors (Mff, Fis1, MiD49, MiD51) regulate mitochondrial fission. The dynamic nature of the mitochondrial network has become evident and detectable, thanks to recent advances in live imaging video microscopy and to the availability of mitochondria-tagged fluorescent proteins. High-resolution confocal reconstruction of mitochondria over time allows researchers to visualize mitochondria shape changes in living cells, under different experimental conditions. Moreover, in recent years, different techniques in living cells have been developed to study the process of mitochondria fusion in more details. Among them are fluorescence recovery after photobleaching (FRAP) of mitochondria-tagged GFP (mtGFP), use of photoactivatable mtGFP, polyethylene glycol (PEG)-based fusion of mtGFP and mtRFP cells, and Renilla luciferase assay (for population studies). In addition, in combination with imaging, the analysis of the expression levels of the different mitochondria-shaping proteins, along with that of their activation status, represents a powerful tool to investigate potential modulations of the mitochondrial network. Here, we review this aspect and then mention a number of techniques, with particular attention to their relative protocols.

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.

Measuring transcription dynamics in living cells using a photobleaching approach.

The transcriptional kinetics of RNA polymerase II, the enzyme responsible for mRNA transcription in the nucleoplasm, can be modulated by a variety of factors. It is therefore important to establish experimental systems that will enable the readout of transcription kinetics of specific genes as they occur in real time within individual cells. This can be performed by implementing fluorescent tagging of the mRNA under live-cell conditions. This chapter describes how to generate fluorescently tagged genes and mRNA, and how a photobleaching approach can produce information on mRNA transcription kinetics.

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.

Single-point single-molecule FRAP distinguishes inner and outer nuclear membrane protein distribution.

The normal distribution of nuclear envelope transmembrane proteins (NETs) is disrupted in several human diseases. NETs are synthesized on the endoplasmic reticulum and then transported from the outer nuclear membrane (ONM) to the inner nuclear membrane (INM). Quantitative determination of the distribution of NETs on the ONM and INM is limited in available approaches, which moreover provide no information about translocation rates in the two membranes. Here we demonstrate a single-point single-molecule FRAP microscopy technique that enables determination of distribution and translocation rates for NETs in vivo.

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.

FRAP, FLIM, and FRET: Detection and analysis of cellular dynamics on a molecular scale using fluorescence microscopy.

The combination of fluorescent-probe technology plus modern optical microscopes allows investigators to monitor dynamic events in living cells with exquisite temporal and spatial resolution. Fluorescence recovery after photobleaching (FRAP), for example, has long been used to monitor molecular dynamics both within cells and on cellular surfaces. Although bound by the diffraction limit imposed on all optical microscopes, the combination of digital cameras and the application of fluorescence intensity information on large-pixel arrays have allowed such dynamic information to be monitored and quantified. Fluorescence lifetime imaging microscopy (FLIM), on the other hand, utilizes the information from an ensemble of fluorophores to probe changes in the local environment. Using either fluorescence-intensity or lifetime approaches, fluorescence resonance energy transfer (FRET) microscopy provides information about molecular interactions, with Ångstrom resolution. In this review, we summarize the theoretical framework underlying these methods and illustrate their utility in addressing important problems in reproductive and developmental systems.

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.

Molecular diffusion and binding analyzed with FRAP.

Intracellular molecular transport and localization are crucial for cells (plant cells as much as mammalian cells) to proliferate and to adapt to diverse environmental conditions. Here, some aspects of the microscopy-based method of fluorescence recovery after photobleaching (FRAP) are introduced. In the course of the last years, this has become a very powerful tool to study dynamic processes in living cells and tissue, and it is expected to experience further increasing demand because quantitative information on biological systems becomes more and more important. This review introduces the FRAP methodology, including some theoretical background, describes challenges and pitfalls, and presents some recent advanced applications.

Monitoring and quantifying dynamic physiological processes in live neurons using fluorescence recovery after photobleaching.

The direct visualization of subcellular dynamic processes is often hampered by limitations in the resolving power achievable with conventional microscopy techniques. Fluorescence recovery after photobleaching has emerged as a highly informative approach to address this challenge, permitting the quantitative measurement of the movement of small organelles and proteins in living functioning cells, and offering detailed insights into fundamental cellular phenomena of physiological importance. In recent years, its implementation has benefited from the increasing availability of confocal microscopy systems and of powerful labeling techniques based on genetically encoded fluorescent proteins or other chemical markers. In this review, we present fluorescence recovery after photobleaching and related techniques in the context of contemporary neurobiological research and discuss quantitative and semi-quantitative approaches to their interpretation.

Fluorescence recovery after photobleaching.

This chapter describes the use of microscope-based fluorescence recovery after photobleaching (FRAP). To quantify the dynamics of proteins within a subcellular compartment, we first outline the general aspects of FRAP experiments and then provide a detailed protocol of how to measure and analyse the most important parameters of FRAP experiments such as mobile fraction and half-time of recovery.

Lessons in fluctuation correlation spectroscopy.

Molecular diffusion and transport processes are fundamental in physical, chemical, and biological systems. Current approaches to measuring molecular transport in cells and tissues based on perturbation methods, e.g., fluorescence recovery after photobleaching, are invasive; single-point fluctuation correlation methods are local; and single-particle tracking requires the observation of isolated particles for relatively long periods of time. We discuss here the detection of molecular transport by exploiting spatiotemporal correlations measured among points at large distances (>1 µm). We illustrate the evolution of the conceptual framework that started with single-point fluorescence fluctuation analysis based on the transit of fluorescent molecules through a small volume of illumination. This idea has evolved to include the measurement of fluctuations at many locations in the sample using microscopy imaging methods. Image fluctuation analysis has become a rich and powerful technique that can be used to extract information about the spatial distribution of molecular concentration and transport in cells and tissues.

Measuring diffusion coefficients via two-photon fluorescence recovery after photobleaching.

Multi-fluorescence recovery after photobleaching is a microscopy technique used to measure the diffusion coefficient (or analogous transport parameters) of macromolecules, and can be applied to both in vitro and in vivo biological systems. Multi-fluorescence recovery after photobleaching is performed by photobleaching a region of interest within a fluorescent sample using an intense laser flash, then attenuating the beam and monitoring the fluorescence as still-fluorescent molecules from outside the region of interest diffuse in to replace the photobleached molecules. We will begin our demonstration by aligning the laser beam through the Pockels Cell (laser modulator) and along the optical path through the laser scan box and objective lens to the sample. For simplicity, we will use a sample of aqueous fluorescent dye. We will then determine the proper experimental parameters for our sample including, monitor and bleaching powers, bleach duration, bin widths (for photon counting), and fluorescence recovery time. Next, we will describe the procedure for taking recovery curves, a process that can be largely automated via LabVIEW (National Instruments, Austin, TX) for enhanced throughput. Finally, the diffusion coefficient is determined by fitting the recovery data to the appropriate mathematical model using a least-squares fitting algorithm, readily programmable using software such as MATLAB (The Mathworks, Natick, MA).

Recent applications of fluorescence recovery after photobleaching (FRAP) to membrane bio-macromolecules.

This review examines some recent applications of fluorescence recovery after photobleaching (FRAP) to biopolymers, while mainly focusing on membrane protein studies. Initially, we discuss the lateral diffusion of membrane proteins, as measured by FRAP. Then, we talk about the use of FRAP to probe interactions between membrane proteins by obtaining fundamental information such as geometry and stoichiometry of the interacting complex. Afterwards, we discuss some applications of FRAP at the cellular level as well as the level of organisms. We conclude by comparing diffusion coefficients obtained by FRAP and several other alternative methods.

Overview of laser microbeam applications as related to antibody targeting.

This chapter reviews several techniques which combine the use of laser microbeams with antibodies to study molecular and cellular biology. An overview of the basic properties of lasers and their integration with microscopes and computers is provided. Biophysical applications, such as fluorescence recovery after photobleaching to measure molecular mobility and fluorescence resonance energy transfer to measure molecular distances, as well as ablative applications for the selective inactivation of proteins or the selective killing of cells are described. Other techniques, such as optical trapping, that do not rely on the interaction of the laser with the targeting antibody, are also discussed.

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.

Line FRAP with the confocal laser scanning microscope for diffusion measurements in small regions of 3-D samples.

We present a truly quantitative fluorescence recovery after photobleaching (FRAP) model for use with the confocal laser scanning microscope based on the photobleaching of a long line segment. The line FRAP method is developed to complement the disk FRAP method reported before. Although being more subject to the influence of noise, the line FRAP model has the advantage of a smaller bleach region, thus allowing for faster and more localized measurements of the diffusion coefficient and mobile fraction. The line FRAP model is also very well suited to examine directly the influence of the bleaching power on the effective bleaching resolution. We present the outline of the mathematical derivation, leading to a final analytical expression to calculate the fluorescence recovery. We examine the influence of the confocal aperture and the bleaching power on the measured diffusion coefficient to find the optimal experimental conditions for the line FRAP method. This will be done for R-phycoerythrin and FITC-dextrans of various molecular weights. The ability of the line FRAP method to measure correctly absolute diffusion coefficients in three-dimensional samples will be evaluated as well. Finally we show the application of the method to the simultaneous measurement of free green fluorescent protein diffusion in the cytoplasm and nucleus of living A549 cells.

Time-resolved Hadamard fluorescence imaging.

We present a new concept for fluorescence lifetime imaging (FLIM) based on time-resolved Hadamard imaging (HI). HI allows image acquisition by use of one single-point detector without requiring a moving scanning stage. Moreover, it reduces the influence of detector noise compared with raster scanning. By use of Monte Carlo simulations it could be confirmed that Hadamard transformation may decrease the error in lifetime estimation and in general in fluorescence parameter estimation when the signal-to-noise ratio is low and detector dark noise is high. This concept may find applications whenever the performance of FLIM or similar methods is limited by high dark-count rates and when the use of a single-point detector is preferable.

Fluorescence recovery after photobleaching: application to nuclear proteins.

Fluorescence redistribution after photobleaching (FRAP) has received increasing attention ever since it was first introduced into cell biological research. The method was developed in the 1970s, when its biological application mainly focused on the mobility of fluorescently labelled constituents of the cell membrane. The development of confocal scanning microscopy in the 1980s facilitated accurate investigation of the behaviour of molecules in the inside of cells without specialised equipment. However, FRAP did not yet become as popular as it is today, probably because of the dedicated and time-consuming methodology required to purify and label proteins or other compounds and, moreover, to inject them into cells. The revolution created by the development of GFP-technology finally lead to a tremendous boost of FRAP applications in studying the behaviour of proteins in the living cells. Finally, the ongoing increase of speed and memory of personal computers allows computer modelling of FRAP experiments for analysis of complex 3-D FRAP data, and for the development of new FRAP assays. Here we discuss several variants of FRAP on the basis of its application to the investigation of the behaviour of proteins in the living cell nucleus.