High-contrast imaging of fluorescent protein FRET by fluorescence polarization microscopy

Detection of Forster resonance energy transfer (FRET) between fluorescent protein labeled targets is a valuable strategy for measurement of protein-protein interactions and other intracellular processes. Despite the utility of FRET, widespread application of this technique to biological problems and high-throughput screening has been limited by low-contrast measurement strategies that rely on the detection of sensitized emission or photodestruction of the sample. Here we report a FRET detection strategy based on detecting depolarized sensitized emission. In the absence of FRET, we show that fluorescence emission from a donor fluorescent protein is highly polarized. Depolarization of fluorescence emission is observed only in the presence of energy transfer. A simple detection strategy was adapted for fluorescence microscopy using both laser scanning and wide-field approaches. This approach is able to distinguish FRET between linked and unlinked Cerulean and Venus fluorescent proteins in living cells with a larger dynamic range than other approaches.     

Quantitative imaging of protein interactions in the cell nucleus

Over the past decade, genetically encoded fluorescent proteins have become widely used as noninvasive markers in living cells. The development of fluorescent proteins, coupled with advances in digital imaging, has led to the rapid evolution of live-cell imaging methods. These approaches are being applied to address biological questions of the recruitment, co-localization, and interactions of specific proteins within particular subcellular compartments. In the wake of this rapid progress, however, come important issues associated with the acquisition and analysis of ever larger and more complex digital imaging data sets. Using protein localization in the mammalian cell nucleus as an example, we will review some recent developments in the application of quantitative imaging to analyze subcellular distribution and co-localization of proteins in populations of living cells. In this report, we review the principles of acquiring fluorescence resonance energy transfer (FRET) microscopy measurements to define the spatial relationships between proteins. We then discuss how fluorescence lifetime imaging microscopy (FLIM) provides a method that is independent of intensity-based measurements to detect localized protein interactions with spatial resolution. Finally, we consider potential problems associated with the expression of proteins fused to fluorescent proteins for FRET-based measurements from living cells.     

Imaging molecular interactions in living cells

Hormones integrate the activities of their target cells through receptor-modulated cascades of protein interactions that ultimately lead to changes in cellular function. Understanding how the cell assembles these signaling protein complexes is critically important to unraveling disease processes, and to the design of therapeutic strategies. Recent advances in live-cell imaging technologies, combined with the use of genetically encoded fluorescent proteins, now allow the assembly of these signaling protein complexes to be tracked within the organized microenvironment of the living cell. Here, we review some of the recent developments in the application of imaging techniques to measure the dynamic behavior, colocalization, and spatial relationships between proteins in living cells. Where possible, we discuss the application of these different approaches in the context of hormone regulation of nuclear receptor localization, mobility, and interactions in different subcellular compartments. We discuss measurements that define the spatial relationships and dynamics between proteins in living cells including fluorescence colocalization, fluorescence recovery after photobleaching, fluorescence correlation spectroscopy, fluorescence resonance energy transfer microscopy, and fluorescence lifetime imaging microscopy. These live-cell imaging tools provide an important complement to biochemical and structural biology studies, extending the analysis of protein-protein interactions, protein conformational changes, and the behavior of signaling molecules to their natural environment within the intact cell.     

Three-dimensional FRET reconstruction microscopy for analysis of dynamic molecular interactions in live cells

Analysis of cellular pathways requires concentration measurements of dynamically interacting molecules within the three-dimensional (3D) space of single living cells. Förster resonance energy transfer (FRET) microscopy from widefield, from confocal, and potentially from superresolution microscopes can access this information; however, these measurements are distorted by the inherent 3D blurring of optical imaging, spectral overlap of fluorophores, and detection noise. We propose a mathematical model of these processes and demonstrate, through simulation, how these distortions limit the dynamic range and sensitivity of conventional FRET microscopy. Using this model, we devise and validate a new approach (called 3D-FRET stoichiometry reconstruction, 3DFSR) for reconstructing 3D distributions of bound and free fluorescent molecules. Previous attempts to reconstruct 3D-FRET data relied on sequential spectral unmixing and deconvolution, a process that corrupts the detection statistics. We demonstrate that 3DFSR is superior to these approaches since it simultaneously models spectral mixing, optical blurring, and detection noise. To achieve the full potential of this technique, we developed an instrument capable of acquiring 3D-FRET data rapidly and sensitively from single living cells. Compared with conventional FRET microscopy, our 3D-FRET reconstruction technique and new instrumentation provides orders of magnitude gains in both sensitivity and accuracy wherein sustained high-resolution four-dimensional (x,y,z,t) imaging of molecular interactions inside living cells was achieved. These results verify previous observations that Cdc42 signaling is localized to the advancing margins of forming phagosomes in macrophages.     

Visualization of ternary complexes in living cells by using a BiFC-based FRET assay

Studies of protein interactions have increased our understanding and knowledge of biological processes. Assays that utilize fluorescent proteins, such as fluorescence resonance energy transfer (FRET) and bimolecular fluorescence complementation (BiFC), have enabled direct visualization of protein interactions in living cells. However, these assays are primarily suitable for a pair of interacting proteins, and methods to visualize and identify multiple protein complexes in vivo are very limited. This protocol describes the recently developed BiFC-FRET assay, which allows visualization of ternary complexes in living cells. We discuss how to design the BiFC-FRET assay on the basis of the validation of BiFC and FRET assays and how to perform transfection experiments for acquisition of fluorescent images for net FRET calculation. We also provide three methods for normalization of the FRET efficiency. The assay employs a two-chromophore and three-filter FRET setup and is applicable to epifluorescence microscopes. The entire protocol takes about 2-3 weeks to complete.     

FRET-FLIM applications in plant systems

A hallmark of cellular processes is the spatio-temporally regulated interplay of biochemical components. Assessing spatial information of molecular interactions within living cells is difficult using traditional biochemical methods. Developments in green fluorescent protein technology in combination with advances in fluorescence microscopy have revolutionised this field of research by providing the genetic tools to investigate the spatio-temporal dynamics of biomolecules in live cells. In particular, fluorescence lifetime imaging microscopy (FLIM) has become an inevitable technique for spatially resolving cellular processes and physical interactions of cellular components in real time based on the detection of Förster resonance energy transfer (FRET). In this review, we provide a theoretical background of FLIM as well as FRET-FLIM analysis. Furthermore, we show two cases in which advanced microscopy applications revealed many new insights of cellular processes in living plant cells as well as in whole plants.     

FRET and FRAP imaging: Approaches to characterise tau and stathmin interactions with microtubules in cells

Microtubules (MTs) are involved in many crucial processes such as cell morphogenesis, mitosis and motility. These dynamic structures resulting from the complex assembly of tubulin are tightly regulated by stabilising MT‐associated proteins (MAPs) such as tau and destabilising proteins, notably stathmin. Because of their key role, these MAPs and their interactions have been extensively studied using biochemical and biophysical approaches, particularly in vitro. Nevertheless, numerous questions remain unanswered and the mechanisms of interaction between MT and these proteins are still unclear in cells. Techniques coupling cell imaging and fluorescence methods, such as Förster resonance energy transfer and fluorescence recovery after photobleaching, are excellent tools to study these interactions in situ. After describing these methods, we will present emblematic data from the literature and unpublished experimental results from our laboratory concerning the interactions between MTs, tau and stathmin in cells.     

Reliable and global measurement of fluorescence resonance energy transfer using fluorescence microscopes

Green fluorescence protein (GFP)-based fluorescence resonance energy transfer (FRET) is increasingly used in investigation of inter- and intramolecular interactions in living cells. In this report, we present a modified method for FRET quantification in cultured cells using conventional fluorescence microscopy. To reliably measure FRET, three positive control constructs in which a cyan fluorescence protein and a yellow fluorescence protein were linked by peptides of 15, 24, or 37 amino acid residues were prepared. FRET was detected using a spectrofluorometer, a laser scanning confocal microscope, and an inverted fluorescence microscope. Three calculation methods for FRET quantification using fluorescence microscopes were compared. By normalization against expression levels of GFP fusion proteins, the modified method gave consistent FRET values that could be compared among different cells with varying protein expression levels. Whole-cell global analysis using this method allowed FRET measurement with high spatial resolutions. Using such a procedure, the interaction of synaptic proteins syntaxin and the synaptosomal associated protein of 25 kDa (SNAP-25) was examined in PC12 cells, which showed strong FRET on plasma membranes. These results demonstrate the effectiveness of the modified method for FRET measurement in live cell systems.     

Imaging protein-protein interactions using fluorescence resonance energy transfer microscopy

Fluorescence resonance energy transfer (FRET) detects the proximity of fluorescently labeled molecules over distances >100 A. When performed in a fluorescence microscope, FRET can be used to map protein-protein interactions in vivo. We here describe a FRET microscopy method that can be used to determine whether proteins that are colocalized at the level of light microscopy interact with one another. This method can be implemented using digital microscopy systems such as a confocal microscope or a wide-field fluorescence microscope coupled to a charge-coupled device (CCD) camera. It is readily applied to samples prepared with standard immunofluorescence techniques using antibodies labeled with fluorescent dyes that act as a donor and acceptor pair for FRET. Energy transfer efficiencies are quantified based on the release of quenching of donor fluorescence due to FRET, measured by comparing the intensity of donor fluorescence before and after complete photobleaching of the acceptor. As described, this method uses Cy3 and Cy5 as the donor and acceptor fluorophores, but can be adapted for other FRET pairs including cyan fluorescent protein and yellow fluorescent protein.     

Fluorescent protein FRET: the good, the bad and the ugly

Dynamic protein interactions play a significant part in many cellular processes. A technique that shows considerable promise in elucidating such interactions is F?rster resonance energy transfer (FRET). When combined with multiple, colored fluorescent proteins, FRET permits high spatial resolution assays of protein-protein interactions in living cells. Because FRET signals are usually small, however, their measurement requires careful interpretation and several control experiments. Nevertheless, the use of FRET in cell biological experiments has exploded over the past few years. Here we describe the physical basis of FRET and the fluorescent proteins appropriate for these experiments. We also review the approaches that can be used to measure FRET, with particular emphasis on the potential artifacts associated with each approach.     

Imaging Protein-protein Interactions in vivo

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, preventing their identification and analysis through traditional biochemical methods such as co-immunoprecipitation. In this regard, the genetically encodable fluorescent proteins (GFP, RFP, etc.) and their associated overlapping fluorescence spectrum have revolutionized our ability to monitor weak interactions in vivo using Förster resonance energy transfer (FRET)^1-3. Here, we detail our use of a FRET-based proximity assay for monitoring receptor-receptor interactions on the endothelial cell surface.     

Imaging protein-protein interactions in living cells

The complex organization of plant cells makes it likely that the molecular behaviour of proteins in the test tube and the cell is different. For this reason, it is essential though a challenge to study proteins in their natural environment. Several innovative microspectroscopic approaches provide such possibilities, combining the high spatial resolution of microscopy with spectroscopic techniques to obtain information about the dynamical behaviour of molecules. Methods to visualize interaction can be based on FRET (fluorescence detected resonance energy transfer), for example in fluorescence lifetime imaging microscopy (FLIM). Another method is based on fluorescence correlation spectroscopy (FCS) by which the diffusion rate of single molecules can be determined, giving insight into whether a protein is part of a larger complex or not. Here, both FRET- and FCS-based approaches to study protein-protein interactions in vivo are reviewed.     

Differences in the mechanisms of proapoptotic BH3 proteins binding to Bcl-XL and Bcl-2 quantified in live MCF-7 cells

Overexpression of antiapoptotic proteins including Bcl-XL and/or Bcl-2 contributes to tumor initiation, progression, and resistance to therapy by direct interactions with proapoptotic BH3 proteins. Release of BH3 proteins from antiapoptotic proteins kills some cancer cells and sensitizes others to chemotherapy. Binding of Bcl-XL and Bcl-2 to the BH3 proteins Bad, Bid, and the three major isoforms of Bim was measured for fluorescent protein fusions in live cells using fluorescence lifetime imaging microscopy and fluorescence resonance energy transfer. In cells the binding of the proteins at mitochondria is similar to the results from in vitro measurements. However, mutations in the BH3 region of Bim known to inhibit binding to Bcl-XL and Bcl-2 in vitro had much less effect in MCF-7 cells. Moreover, the BH3 mimetic ABT-737 inhibited Bad and Bid but not Bim binding to Bcl-XL and Bcl-2. Thus, the selectivity of ABT-737 also differs markedly from predictions made from in vitro measurements.     

Cytology of fungal pathogens and plant-host interactions

Imaging plays a unique role in fungal cell biology and phytopathology by allowing for the documentation of molecular structure in individual fixed and living cells. Advances in fluorescence laser techniques, including confocal and multiphoton microscopy, are opening new avenues for cellular exploration. These techniques hold tremendous potential for studies of host-pathogen interactions including the use of genetically encoded markers such as green fluorescent protein, in situ hybridization and fluorescence resonance energy transfer.     

Monitoring receptor-mediated activation of heterotrimeric G-proteins by fluorescence resonance energy transfer

Green fluorescent protein (GFP)-centered fluorescence resonance energy transfer (FRET) relies on a distance-dependent transfer of energy from a donor fluorophore to an acceptor fluorophore and can be used to examine protein interactions in living cells. Here we describe a method to monitor the association and disassociation of heterotrimeric GTP-binding (G-proteins) from one another before and after stimulation of coupled receptors in living Dictyostelium discoideum cells. The Galpha(2)and Gbetagamma proteins were tagged with cyan and yellow fluorescent proteins and used to observe the state of the G-protein heterotrimer. Data from emission spectra were used to detect the FRET fluorescence and to determine kinetics and dose-response curves of bound ligand and analogs. Extending G-protein FRET to mammalian G-proteins should enable direct in situ mechanistic studies and applications such as drug screening and identifying ligands of new G-protein-coupled receptors.