Fluorescence resonance energy transfer of GFP and YFP by spectral imaging and quantitative acceptor photobleaching

To study protein–protein interactions by fluorescence energy transfer (FRET), the proteins of interest are tagged with either a donor or an acceptor fluorophore. For efficient FRET, fluorophores need to have a reasonable overlap of donor emission and acceptor excitation spectra. However, given the relatively small Stokes shift of conventional fluorescent proteins, donor and acceptor pairs with high FRET efficiencies have emission spectra that are difficult to separate. GFP and YFP are widely used in fluorescence microscopy studies. The spectral qualities of GFP and YFP make them one of the most efficient FRET donor–acceptor couples available. However, the emission peaks of GFP (510 nm) and YFP (527 nm) are spectrally too close for separation by conventional fluorescence microscopy. Difficulties in simultaneous detection of GFP and YFP with a fluorescence microscope are eliminated when spectral imaging and subsequent linear unmixing are applied. This allows FRET microscopy using these tags to study protein–protein interactions. We adapted the linear unmixing procedure from commercially available software (Zeiss) for use with acceptor photobleaching FRET using GFP and YFP as FRET pair. FRET efficiencies up to 52% for a GFP-YFP fusion protein were measured. To investigate the applicability of the procedure, we used two constituents of the nucleotide excision repair system, which removes UV-induced single-strand DNA damage. ERCC1 and XPF form a heterodimeric 5' endonuclease in nucleotide excision repair. FRET between ERCC1-GFP and XPF-YFP occurs with an efficiency of 30%.     

Quantitative fluorescence resonance energy transfer (FRET) measurement with acceptor photobleaching and spectral unmixing

Fluorescence resonance energy transfer (FRET) by acceptor photobleaching is a simple but effective tool for measurements of protein–protein interactions. Until recently, it has been restricted to qualitative or relative assessments owing to the spectral bleed‐through contamination resulting from fluorescence overlap between the donor and the acceptor. In this paper, we report a quantitative algorithm that combines the spectral unmixing technique with FRET by acceptor photobleaching. By spectrally unmixing the emissions before and after photobleaching, it is possible to resolve the spectral bleed‐through and retrieve the FRET efficiency/interaction distance quantitatively. Using a human keratinocyte cell line transfected with cyan fluorescent protein (CFP)‐ and yellow fluorescent protein (YFP)‐tagged Cx26 connexins as an example, FRET information at homotypic gap junctions is measured and compared with well‐established methods. Results indicate that the new approach is sensitive, flexible, instrument independent and solely FRET dependent. It can achieve FRET estimations similar to that from a sensitized emission FRET method. This approach has a great advantage in providing the relative concentrations of the donor and the acceptor; this is, for example, very important in the comparative study of cell populations with variable expression levels.     

FRET and colocalization analyzer--a method to validate measurements of sensitized emission FRET acquired by confocal microscopy and available as an ImageJ Plug-in

Fluorescence resonance energy transfer (FRET) between an adequate pair of fluorophores is an indication of closer proximity than colocalization and is used by biologists to study fluorescently modified protein interactions inside cells. We present a method for visualization of FRET images acquired by confocal sensitized emission, involving excitation of the donor fluorophore and detection of the energy transfer as an emission from the acceptor fluorophore into the FRET channel. Authentic FRET signal measurements require the correction from the FRET channel of the undesired bleed-through signals (BT) resulting from both the leak-through of the donor emission and the direct acceptor emission. Our method reduces the interference of the user to a minimum by analyzing the entire image, pixel by pixel. It proposes imaging treatments and the display of control images to validate the BT calculation and the image corrections. It displays FRET images as a function of the colocalization of the two fluorescent partners. Finally, it proposes an alternative to normalization of the FRET intensities to compare FRET signal variations between samples. This method called "FRET and Colocalization Analyzer" has been implemented in a Plug-in of the freely available ImageJ software. It is particularly adapted when transient expression of the fluorescent proteins is used thereby giving very variable expression levels or when the colocalization of the two partners is varying in proportion, in amount, and in size, as a function of time. The method and program are validated using the analysis of the spatio-temporal interactions between a G-protein coupled receptor, the tachykinin NK2 receptor, and the beta-arrestin 2 as an example.     

Novel lambda FRET spectral confocal microscopy imaging method

We report a highly specific, sensitive, and robust method for analyzing fluorescence resonance energy transfer (FRET) based on spectral laser scanning confocal microscopy imaging. The lambda FRET (lambdaFRET) algorithm comprises imaging of a FRET sample at multiple emission wavelengths rendering a FRET spectrum, which is separated into its donor and acceptor components to obtain a pixel-based calculation of FRET efficiency. The method uses a novel off-line precalibration procedure for spectral bleed-through correction based on the acquisition of reference reflection images, which simplifies the method and reduces variability. LambdaFRET method was validated using structurally characterized FRET standards with variable linker lengths and stoichiometries designed for this purpose. LambdaFRET performed better than other well-established methods, such as acceptor photobleaching and sensitized emission-based methods, in terms of specificity, reproducibility, and sensitivity to distance variations. Moreover, lambdaFRET analysis was unaffected by high fluorochrome spectral overlap and cellular autofluorescence. The lambdaFRET method demonstrated outstanding performance in intra- and intermolecular FRET analysis in both fixed and live cell imaging studies.     

Effects of donor and acceptor's fluorescence lifetimes on the method of applying Förster resonance energy transfer in STED microscopy

Förster resonance energy transfer (FRET) probes being used to improve the resolution of stimulated emission depletion (STED) microscopy are numerically discussed. Besides the FRET efficiency and the excitation intensity, the fluorescence lifetimes of donor and acceptor are found to be another key parameter for the resolution enhancement. Using samples of FRET pairs with shorter donor lifetime and longer acceptor lifetime enhances the nonlinearity of the donor fluorescence, which leads to an increased resolution. The numerical simulation shows that a double resolution improvement of STED microscopy can be achieved by using Cy3–Atto647N samples when compared with that of using standard Cy3‐only samples.     

Multichannel wide‐field microscopic FRET imaging based on simultaneous spectral unmixing of excitation and emission spectra

Simultaneous spectral unmixing of excitation and emission spectra (ExEm unmixing) has inherent ability resolving spectral crosstalks, two key issues of quantitative fluorescence resonance energy transfer (FRET) measurement, of both the excitation and emission spectra between donor and acceptor without additional corrections. We here set up a filter‐based multichannel wide‐field microscope for ExEm unmixing‐based FRET imaging (m‐ExEm‐spFRET) containing a constant system correction factor (f_sc) for a stable system. We performed m‐ExEm‐spFRET with four‐ and two‐wavelength excitation respectively on our system to quantitatively image single living cells expressing FRET tandem constructs, and obtained accurate FRET efficiency (E) and concentration ratio of acceptor to donor (R_C). We also performed m‐ExEm‐spFRET imaging for single living cells coexpressing CFP‐Bax and YFP‐Bax, and found that the E values were about 0 for control cells and about 28% for staurosporin‐treated cells when R_C were larger than 1, indicating that staurosporin induced significant oligomerisation.     

Spectral resolution in conjunction with polar plots improves the accuracy and reliability of FLIM measurements and estimates of FRET efficiency

A spectrograph with continuous wavelength resolution has been integrated into a frequency‐domain fluorescence lifetime‐resolved imaging microscope (FLIM). The spectral information assists in the separation of multiple lifetime components, and helps resolve signal cross‐talking that can interfere with an accurate analysis of multiple lifetime processes. This extends the number of different dyes that can be measured simultaneously in a FLIM measurement. Spectrally resolved FLIM (spectral‐FLIM) also provides a means to measure more accurately the lifetime of a dim fluorescence component (as low as 2% of the total intensity) in the presence of another fluorescence component with a much higher intensity. A more reliable separation of the donor and acceptor fluorescence signals are possible for Förster resonance energy transfer (FRET) measurements; this allows more accurate determinations of both donor and acceptor lifetimes. By combining the polar plot analysis with spectral‐FLIM data, the spectral dispersion of the acceptor signal can be used to derive the donor lifetime – and thereby the FRET efficiency – without iterative fitting. The lifetime relation between the donor and acceptor, in conjunction with spectral dispersion, is also used to separate the FRET pair signals from the donor alone signal. This method can be applied further to quantify the signals from separate FRET pairs, and provide information on the dynamics of the FRET pair between different states.     

Fluorescence resonance energy transfer (FRET) measurement by gradual acceptor photobleaching

Fluorescence resonance energy transfer (FRET) is an extremely effective tool to detect molecular interaction at suboptical resolutions. One of the techniques for measuring FRET is acceptor photobleaching: the increase in donor fluorescence after complete acceptor photobleaching is a measure of the FRET efficiency. However, in wide-field microscopy, complete acceptor photobleaching is difficult due to the low excitation intensities. In addition, the method is sensitive to inadvertent donor bleaching, autofluorescence and bleed-through of excitation light. In the method introduced in this paper, donor and acceptor intensities are monitored continuously during acceptor photobleaching. Subsequently, curve fitting is used to determine the FRET efficiency. The method was demonstrated on cameleon (YC2.1), a FRET-based Ca²⁺ indicator, and on a CFP-YFP fusion protein expressed in HeLa cells. FRET efficiency of cameleon in the presence of 1 mm Ca²⁺ was 31 ± 3%. In the absence of Ca²⁺ a FRET efficiency of 15 ± 2% was found. A FRET efficiency of 28% was found for the CFP-YFP fusion protein in HeLa cells. Advantages of the method are that it does not require complete acceptor photobleaching, it includes correction for spectral cross-talk, donor photobleaching and autofluorescence, and is relatively simple to use on a normal wide-field microscope.     

Characterization of two-photon excitation fluorescence lifetime imaging microscopy for protein localization

Two-photon excitation fluorescence resonance energy transfer (2P-FRET) imaging microscopy can provide details of specific protein molecule interactions inside living cells. Fluorophore molecules used for 2P-FRET imaging have characteristic absorption and emission spectra that introduce spectral cross-talk (bleed-through) in the FRET signal that should be removed in the 2P-FRET images, to establish that FRET has actually occurred and to have a basis for distance estimations. These contaminations in the FRET signal can be corrected using a mathematical algorithm to extract the true FRET signal. Another approach is 2P-FRET fluorescence lifetime imaging (FLIM). This methodology allows studying the dynamic behavior of protein-protein interactions in living cells and tissues. 2P-FRET-FLIM was used to study the dimerization of the CAATT/enhancer binding protein alpha (C/EBPalpha). Results show that the reduction in donor lifetime in the presence of acceptor reveals the dimerization of the protein molecules and also determines more precisely the distance between the donor and acceptor. We describe the development and characterization of the 2P-FRET-FLIM imaging system with the Bio-Rad Radiance2100 confocal/multiphoton microscopy system.     

Design and analysis of multi-color confocal microscopy with a wavelength scanning detector

Spectral (or multi-color) microscopy has the ability to detect the fluorescent light of biological specimens with a broad range of wavelengths. Currently, the acousto-optic tunable filter (AOTF) is widely used in spectral microscopy as a substitute for a multiple-dichroic mirror to divide excitation and emission signals while maintaining sufficient light efficiency. In addition, systems which utilize an AOTF have a very fast switching speed and high resolution for wavelength selection. In this paper, confocal-spectral microscopy is proposed with a particular spectrometer design with a wavelength-scanning galvano-mirror. This enables the detection of broadband (480-700 nm) fluorescence signals by a single point detector (photomultiplier tube) instead of a CCD pixel array. For this purpose, a number of optical elements were applicably designed. A prism is used to amplify the dispersion angle, and the design of the relay optics matches the signals to the diameter of the wavelength-scanning galvano-mirror. Also, a birefringent material known as calcite is used to offset the displacement error at the image plane depending on the polarization states. The proposed multi-color confocal microscopy with the unique detection body has many advantages in comparison with commercial devices. In terms of the detection method, it can be easily applied to other imaging modalities.     

Quantitative FRET measurement using emission‐spectral unmixing with independent excitation crosstalk correction

Quantification of fluorescence resonance energy transfer (FRET) needs at least two external samples, an acceptor‐only reference and a linked FRET reference, to calibrate fluorescence signal. Furthermore, all measurements for references and FRET samples must be performed under the same instrumental conditions. Based on a novel notion to predetermine the molar extinction coefficient ratio (R_C) of acceptor‐to‐donor for the correction of acceptor excitation crosstalk, we present here a robust and independent emission‐spectral unmixing FRET methodology, Iem‐spFRET, which can simultaneously measure the E and R_C of FRET sample without any external references, such that Iem‐spFRET circumvents the rigorous restriction of keeping the same imaging conditions for all FRET experiments and thus can be used for the direct measurement of FRET sample. We validate Iem‐spFRET by measuring the absolute E and R_C values of standard constructs with different acceptor‐to‐donor stoichiometry expressed in living cells. Our results demonstrate that Iem‐spFRET is a simple and powerful tool for real‐time monitoring the dynamic intermolecular interaction within single living cells.     

Fluorescence resonance energy transfer from cyan to yellow fluorescent protein detected by acceptor photobleaching using confocal microscopy and a single laser

One manifestation of fluorescence resonance energy transfer (FRET) is an increase in donor fluorescence after photobleaching the acceptor. Published acceptor‐photobleaching methods for FRET have mainly used wide‐field microscopy. A laser scanning confocal microscope enables faster and targeted bleaching within the field of view, thereby improving speed and accuracy. Here we demonstrate the approach with CFP and YFP, the most versatile fluorescent markers now available for FRET. CFP/YFP FRET imaging has been accomplished with a single laser (argon) available on virtually all laser‐scanning confocal microscopes. Accordingly, we also describe the conditions that we developed for dual imaging of CFP and YFP with the 458 and 514 argon lines. We detect FRET in a CFP/YFP fusion and also between signalling molecules (TNF‐Receptor‐Associated‐Factors or TRAFs) that are known to homo‐ and heterotrimerize. Importantly, we demonstrate that appropriate controls are essential to avoid false positives in FRET by acceptor photobleaching. We use two types of negative control: (a) an internal negative control (non‐bleached areas of the cell) and (b) cells with donor in the absence of the acceptor (CFP only). We find that both types of negative control can yield false FRET. Given this false FRET background, we describe a method for distinguishing true positive signals. In summary, we extensively characterize a simple approach to FRET that should be adaptable to most laser‐scanning confocal microscopes, and demonstrate its feasibility for detecting FRET between several CFP/YFP partners.     

Fluorescence lifetime imaging--techniques and applications

Fluorescence lifetime imaging (FLIM) uses the fact that the fluorescence lifetime of a fluorophore depends on its molecular environment but not on its concentration. Molecular effects in a sample can therefore be investigated independently of the variable, and usually unknown concentration of the fluorophore. There is a variety of technical solutions of lifetime imaging in microscopy. The technical part of this paper focuses on time‐domain FLIM by multidimensional time‐correlated single photon counting, time‐domain FLIM by gated image intensifiers, frequency‐domain FLIM by gain‐modulated image intensifiers, and frequency‐domain FLIM by gain‐modulated photomultipliers. The application part describes the most frequent FLIM applications: Measurement of molecular environment parameters, protein‐interaction measurements by Förster resonance energy transfer (FRET), and measurements of the metabolic state of cells and tissue via their autofluorescence. Measurements of local environment parameters are based on lifetime changes induced by fluorescence quenching or conformation changes of the fluorophores. The advantage over intensity‐based measurements is that no special ratiometric fluorophores are needed. Therefore, a much wider selection of fluorescence markers can be used, and a wider range of cell parameters is accessible. FLIM‐FRET measures the change in the decay function of the FRET donor on interaction with an acceptor. FLIM‐based FRET measurement does not have to cope with problems like donor bleedthrough or directly excited acceptor fluorescence. This relaxes the requirements to the absorption and emission spectra of the donors and acceptors used. Moreover, FLIM‐FRET measurements are able to distinguish interacting and noninteracting fractions of the donor, and thus obtain independent information about distances and interacting and noninteracting protein fractions. This is information not accessible by steady‐state FRET techniques. Autofluorescence FLIM exploits changes in the decay parameters of endogenous fluorophores with the metabolic state of the cells or the tissue. By resolving changes in the binding, conformation, and composition of biologically relevant compounds FLIM delivers information not accessible by steady‐state fluorescence techniques.     

Multi-dimensional fluorescence lifetime and FRET measurements

When and where proteins associate with each other in living cells are key questions in many biological research projects. One way to address these questions is to measure the extent of F?rster resonance energy transfer (FRET) between proteins that have been labeled with appropriate donor and acceptor fluorophores. When both proteins interact, donor and acceptor fluorophores are brought into close vicinity so that the donor can transmit a part of its excitation energy to the acceptor. As a result, both the intensity and the lifetime of the donor fluorescence decrease, whereas the intensity of the acceptor emission increases. This offers different approaches to determine FRET efficiency: One is to detect changes in the intensity of donor and acceptor emission, the other is to measure changes in the lifetime of the donor molecule. One important advantage of the fluorescence lifetime approach is that it allows to distinguish between free and associated donor molecules. However, like intensity measurements it lacks an intrinsic control ensuring that changes in the measured parameters are only due to FRET and not other quenching processes. Here, we show how this limitation can be overcome by spectrally resolved fluorescence lifetime measurements in the time domain. One technique is based on a streak camera system, the other technique is based on a time-correlated-single-photon-counting approach. Both approaches allow biologists to record both donor and acceptor fluorescence emitted by the sample in a single measurement.     

Imaging FRET standards by steady-state fluorescence and lifetime methods

Imaging fluorescence resonance energy transfer (FRET) between molecules labeled with fluorescent proteins is emerging as a powerful tool to study changes in ions, ligands, and molecular interactions in their physiological cellular environment. Different methods use either steady-state fluorescence properties or lifetime to quantify the FRET rate. In addition, some provide the absolute FRET efficiency whereas others are simply a relative index very much influenced by the actual settings and instrumentation used, which makes the interpretation of a given FRET rate very difficult. The use and exchange of FRET standards in laboratories using these techniques would help to overcome this drawback. We report here the construction and systematic evaluation of FRET standard probes of varying FRET efficiencies. The standards for intramolecular FRET were protein fusions of the cyan and yellow variants of A. victoria green fluorescent protein (ECFP and citrine) joined by short linkers or larger protein spacers, or ECFP tagged with a tetracysteine motif and labeled with the biarsenical fluorochrome, FlAsH. Negative and positive controls of intermolecular FRET were also used. We compared these FRET standards with up to four FRET quantification methods: ratioing of acceptor to donor emission, donor intensity recovery upon acceptor photobleach, sensitized emission after spectral unmixing of raw images, and fluorescence lifetime imaging (FLIM). The latter was obtained with a frequency-domain setup able to provide high quality lifetime images in less than a second, and is thus very well suited for live cell studies. The FRET rates or indexes of the standards were in good agreement regardless of the method used. For the CFP-tetraCys/FlAsH pair, the rate calculated from CFP quenching was faster than that obtained by FLIM.     

Probing plasma membrane microdomains in cowpea protoplasts using lipidated GFP-fusion proteins and multimode FRET microscopy

Multimode fluorescence resonance energy transfer (FRET) microscopy was applied to study the plasma membrane organization using different lipidated green fluorescent protein (GFP)‐fusion proteins co‐expressed in cowpea protoplasts. Cyan fluorescent protein (CFP) was fused to the hyper variable region of a small maize GTPase (ROP7) and yellow fluorescent protein (YFP) was fused to the N‐myristoylation motif of the calcium‐dependent protein kinase 1 (LeCPK1) of tomato. Upon co‐expressing in cowpea protoplasts a perfect co‐localization at the plasma membrane of the constructs was observed. Acceptor‐photobleaching FRET microscopy indicated a FRET efficiency of 58% in protoplasts co‐expressing CFP‐Zm7hvr and myrLeCPK1‐YFP, whereas no FRET was apparent in protoplasts co‐expressing CFP‐Zm7hvr and YFP. Fluorescence spectral imaging microscopy (FSPIM) revealed, upon excitation at 435 nm, strong YFP emission in the fluorescence spectra of the protoplasts expressing CFP‐Zm7hvr and myrLeCPK1‐YFP. Also, fluorescence lifetime imaging microscopy (FLIM) analysis indicated FRET because the CFP fluorescence lifetime of CFP‐Zm7hvr was reduced in the presence of myrLeCPK1‐YFP. A FRET fluorescence recovery after photobleaching (FRAP) analysis on a partially acceptor‐bleached protoplast co‐expressing CFP‐Zm7hvr and myrLeCPK1‐YFP revealed slow requenching of the CFP fluorescence in the acceptor‐bleached area upon diffusion of unbleached acceptors into this area. The slow exchange of myrLeCPK1‐YFP in the complex with CFP‐Zm7hvr reflects a relatively high stability of the complex. Together, the FRET data suggest the existence of plasma membrane lipid microdomains in cowpea protoplasts.     

Ensemble and single particle photophysical properties (two-photon excitation, anisotropy, FRET, lifetime, spectral conversion) of commercial quantum dots in solution and in live cells

In this work, we characterized streptavidin-conjugated quantum dots (QDs) manufactured by Quantum Dot Corporation. We present data on: (1) two-photon excitation; (2) fluorescence lifetimes; (3) ensemble and single QD emission anisotropy; (4) QDs as donors for Forster resonance energy transfer (FRET); and (5) spectral conversion of QDs exposed to high-intensity illumination. We also demonstrate the utility of QDs for (1) imaging the binding and uptake of biotinylated transferrin on living cells, and (2) resolving by fluorescence lifetime imaging microscopy (FLIM) signals originating from QDs from those of spatially and spectrally overlapping visible fluorescent proteins (VFPs).     

PixFRET, an ImageJ plug-in for FRET calculation that can accommodate variations in spectral bleed-throughs

Fluorescence resonance energy transfer (FRET) allows the user to investigate interactions between fluorescent partners. One crucial issue when calculating sensitized emission FRET is the correction for spectral bleed-throughs (SBTs), which requires to calculate the ratios between the intensities in the FRET and in the donor or acceptor settings, when only the donor or acceptor are present. Theoretically, SBT ratios should be constant. However, experimentally, these ratios can vary as a function of fluorophore intensity, and assuming constant values may hinder precise FRET calculation. One possible cause for such a variation is the use of a microscope set-up with different photomultipliers for the donor and FRET channels, a set-up allowing higher speed acquisitions on very dynamic fluorescent molecules in living cells. Herein, we show that the bias introduced by the differential response of the two PMTs can be circumvented by a simple modeling of the SBT ratios as a function of fluorophore intensity. Another important issue when performing FRET is the localization of FRET within the cell or a population of cells. We hence developed a freely available ImageJ plug-in, called PixFRET, that allows a simple and rapid determination of SBT parameters and the display of normalized FRET images. The usefulness of this modeling and of the plug-in are exemplified by the study of FRET in a system where two interacting nuclear receptors labeled with ECFP and EYFP are coexpressed in living cells.