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.     

Sensitivity of CFP/YFP and GFP/mCherry pairs to donor photobleaching on FRET determination by fluorescence lifetime imaging microscopy in living cells

Fluorescent protein-based FRET is a powerful method for visualizing protein-protein interactions and biochemical reactions in living cells. It can be difficult, however, to avoid photobleaching when observing fluorescent cells under the microscope, especially those expressing CFP. We compared the sensitivity of two protein-based FRET pairs to light-induced fluorescence changes in the donor, on FRET determination by fluorescence lifetime imaging microscopy (FLIM). Thanks to the very low excitation light levels of the time- and space-correlated single photon counting (TSCSPC) method, FLIM acquisitions were achieved without donor photobleaching. Here, we show that photobleaching of CFP by a mercury lamp under the microscope induced a decrease in the mean fluorescence lifetime, which interfered with FRET determination between CFP and YFP. Importantly, the range of light-induced variation of the mean fluorescence lifetime of CFP was not proportional to the decrease in the steady state fluorescence intensity and varied from cell to cell. The choice of the CFP/YFP pair therefore requires that the cells be observed and analyzed at very low light levels during the whole FRET experiment. In contrast, the GFP/mCherry pair provided an accurate FRET measurement by FLIM, even if some GFP photobleaching took place. We thus demonstrate that CFP can be an unreliable donor for FRET determination in living cells, due to its photosensitivity properties. We demonstrate that the GFP/mCherry pair is better suited for FRET measurement by FLIM in living cells than the CFP/YFP pair.     

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.     

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%.     

Robust approaches to quantitative ratiometric FRET imaging of CFP/YFP fluorophores under confocal microscopy

Ratiometric quantification of CFP/YFP FRET enables live-cell time-series detection of molecular interactions, without the need for acceptor photobleaching or specialized equipment for determining fluorescence lifetime. Although popular in widefield applications, its implementation on a confocal microscope, which would enable sub-cellular resolution, has met with limited success. Here, we characterize sources of optical variability (unique to the confocal context) that diminish the accuracy and reproducibility of ratiometric FRET determination and devise practical remedies. Remarkably, we find that the most popular configuration, which pairs an oil objective with a small pinhole aperture, results in intractable variability that could not be adequately corrected through any calibration procedure. By quantitatively comparing several imaging configurations and calibration procedures, we find that significant improvements can be achieved by combining a water objective and increased pinhole aperture with a uniform-dye calibration procedure. The combination of these methods permitted remarkably consistent quantification of sub-cellular FRET in live cells. Notably, this methodology can be readily implemented on a standard confocal instrument, and the dye calibration procedure yields a time savings over traditional live-cell calibration methods. In all, identification of key technical challenges and practical compensating solutions promise robust sub-cellular ratiometric FRET imaging under confocal microscopy.     

Fluorescence resonance energy transfer and anisotropy reveals both hetero- and homo-energy transfer in the pleckstrin homology-domain and the parathyroid hormone-receptor

We present a method and an apparatus of polarized fluorescence resonance energy transfer (FRET) and anisotropy imaging microscopy done in parallel for improved interpretation of the photophysical interactions. We demonstrate this apparatus to better determine the protein-protein interactions in the pleckstrin homology domain and the conformational changes in the Parathyroid Hormone Receptor, a G-protein coupled receptor, both fused to the cyan and yellow fluorescent proteins for either inter- or intramolecular FRET. In both cases, the expression levels of proteins and also background autofluorescence played a significant role in the depolarization values measured in association with FRET. The system has the sensitivity and low-noise capability of single-fluorophore detection. Using counting procedures from single-molecule methods, control experiments were performed to determine number densities of green fluorescence protein variants CFP and YFP where homo resonance energy transfer can occur. Depolarization values were also determined for flavins, a common molecule of cellular background autofluorescence. From the anisotropy measurements of donor and acceptor, the latter when directly excited or when excited by energy transfer, we find that our instrumentation and method also characterizes crucial effects from homotransfer, polarization specific photobleaching and background molecules.     

Multi-dimensional time-correlated single photon counting (TCSPC) fluorescence lifetime imaging microscopy (FLIM) to detect FRET in cells

We present a novel, multi‐dimensional, time‐correlated single photon counting (TCSPC) technique to perform fluorescence lifetime imaging with a laser‐scanning microscope operated at a pixel dwell‐time in the microsecond range. The unsurpassed temporal accuracy of this approach combined with a high detection efficiency was applied to measure the fluorescent lifetimes of enhanced cyan fluorescent protein (ECFP) in isolation and in tandem with EYFP (enhanced yellow fluorescent protein). This technique enables multi‐exponential decay analysis in a scanning microscope with high intrinsic time resolution, accuracy and counting efficiency, particularly at the low excitation levels required to maintain cell viability and avoid photobleaching. Using a construct encoding the two fluorescent proteins separated by a fixed‐distance amino acid spacer, we were able to measure the fluorescence resonance energy transfer (FRET) efficiency determined by the interchromophore distance. These data revealed that ECFP exhibits complex exponential fluorescence decays under both FRET and non‐FRET conditions, as previously reported. Two approaches to calculate the distance between donor and acceptor from the lifetime delivered values within a 10% error range. To confirm that this method can be used also to quantify intermolecular FRET, we labelled cultured neurones with the styryl dye FM1‐43, quantified the fluorescence lifetime, then quenched its fluorescence using FM4‐64, an efficient energy acceptor for FM1‐43 emission. These experiments confirmed directly for the first time that FRET occurs between these two chromophores, characterized the lifetimes of these probes, determined the interchromophore distance in the plasma membrane and provided high‐resolution two‐dimensional images of lifetime distributions in living neurones.     

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.     

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.     

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.     

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 detected by scanning near-field optical microscopy

Fluorescence resonance energy transfer (FRET) between excited fluorescent donor and acceptor molecules occurs via the F?rster mechanism over a range of 1-10 nm. Because of the strong (sixth power) distance dependence of the signal, FRET has been used to assess the proximity of molecules in biological systems. We used a scanning near-field optical microscope (SNOM) operated in the shared-aperture mode using uncoated glass fibre tips to detect FRET between dye molecules embedded in polyvinyl alcohol films and bound to cell surfaces. FRET was detected by selective photobleaching of donor and acceptor fluorophores. We also present preliminary results on pixel-by-pixel energy transfer efficiency measurements using SNOM.     

Fluorescence resonance energy transfer detected by scanning near-field optical microscopy

Fluorescence resonance energy transfer (FRET) between excited fluorescent donor and acceptor molecules occurs via the Förster mechanism over a range of 1–10 nm. Because of the strong (sixth power) distance dependence of the signal, FRET has been used to assess the proximity of molecules in biological systems. We used a scanning near-field optical microscope (SNOM) operated in the shared-aperture mode using uncoated glass fibre tips to detect FRET between dye molecules embedded in polyvinyl alcohol films and bound to cell surfaces. FRET was detected by selective photobleaching of donor and acceptor fluorophores. We also present preliminary results on pixel-by-pixel energy transfer efficiency measurements using SNOM.     

Fluorescence localization after photobleaching (FLAP): a new method for studying protein dynamics in living cells

FLAP is a new method for localized photo‐labelling and subsequent tracking of specific molecules within living cells. It is simple in principle, easy to implement and has a wide potential application. The molecule to be located carries two fluorophores: one to be photobleached and the other to act as a reference label. Unlike the related methods of fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP), the use of a reference fluorophore permits the distribution of the photo‐labelled molecules themselves to be tracked by simple image differencing. In effect, FLAP is therefore comparable with methods of photoactivation. Its chief advantage over the method of caged fluorescent probes is that it can be used to track chimaeric fluorescent proteins directly expressed by the cells. Although methods are being developed to track fluorescent proteins by direct photoactivation, these still have serious drawbacks. In order to demonstrate FLAP, we have used nuclear microinjection of cDNA fusion constructs of β‐actin with yellow (YFP) and cyan (CFP) fluorescent proteins to follow both the fast relocation dynamics of monomeric (globular) G‐actin and the much slower dynamics of filamentous F‐actin simultaneously in living cells.     

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.     

Fiber laser based two-photon FRET measurement of calmodulin and mCherry-E(0)GFP proteins

The speed and accuracy of Förster resonance energy transfer (FRET) measurements can be improved by rapidly alternating excitation wavelengths between the donor and acceptor fluorophore. We demonstrate FRET efficiency measurements based on a fiber laser and photonic crystal fiber as the source for two‐photon excitation (TPE). This system offers the potential for rapid wavelength switching with the benefits of axial optical sectioning and improved penetration depth provided by TPE. Correction of FRET signals for cross excitation and cross emission was achieved by switching the excitation wavelength with an electrically controlled modulator. Measurement speed was primarily limited by integration times required to measure fluorescence. Using this system, we measured the FRET efficiency of calmodulin labeled with Alexa Fluor 488 and Texas Red dyes. In addition, we measured two‐photon induced FRET in an E⁰GFP‐mCherry protein construct. Results from one‐photon and two‐photon excitation are compared to validate the rapid wavelength switched two‐photon measurements. Microsc. Res. Tech. 75:837–843, 2012. © 2011 Wiley Periodicals, Inc.     

Fluorescence in the tandem scanning microscope

Our studies have shown that the fluorescence mode can be used to good effect in both tandem scanning microscopes (TSM: direct view confocal microscopes) as well as confocal scanning laser microscopes (CSLM). Applications are presented which show that the two great advantages of TSM are real-time viewing and real colour, which allow faster use and interpretation. CSLM are complementary, not competitive, being currently more sophisticated for low-level fluorescence work. This is equally possible with available TSM, but requires further development using CCD cameras and image-processing systems.     

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.     

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.