Elsevier

Methods

Volume 29, Issue 1, January 2003, Pages 58-73
Methods

Characterization of one- and two-photon excitation fluorescence resonance energy transfer microscopy

https://doi.org/10.1016/S1046-2023(02)00283-9Get rights and content

Abstract

Advances in molecular biology provide various methods to define the structure and function of the individual proteins that form the component parts of subcellular structures. The ability to see the dynamic behavior of a specific protein inside the living cell became possible through the application of advanced fluorescence resonance energy transfer (FRET) microscope techniques. The fluorophore molecule used for FRET imaging has a characteristic absorption and emission spectrum that should be considered for characterizing the FRET signal. In this article we describe the system development for the image acquisition for one- and two-photon excitation FRET microscopy. We also describe the precision FRET (PFRET) data analysis algorithm that we developed to remove spectral bleed-through and variation in the fluorophore expression level (or concentration) for the donor and acceptor molecules. The acquired images have been processed using a PFRET algorithm to calculate the energy transfer efficiency and the distance between donor and acceptor molecules. We implemented the software correction to study the organization of the apical endosome in epithelial polarized MDCK cells and dimerization of the CAATT/enhancer binding protein α (C/EBPα). For these proteins, the results revealed that the extent of correction affects the conventionally calculated energy transfer efficiency (E) and the distance (r) between donor and acceptor molecules by 38 and 9%, respectively.

Introduction

Within the living cell, interacting proteins are assembled into molecular machines that function to control cellular homeostasis. These protein assemblies are traditionally studied using biophysical or biochemical methods such as affinity chromatography and coimmunoprecipitation. Recently, two-hybrid and phage-display methods have been used for detecting protein–protein interactions. These in vitro screening methods have the advantage of providing direct access to the genetic information encoding unknown protein partners [1]. These techniques do not allow direct access to interactions of these protein partners in their natural environment inside the living cell, but using the approach of fluorescence resonance energy transfer (FRET) microscopy, this information can be obtained from single living cells with nanometer resolution [2], [3], [4], [5], [6], [7], [8], [9].

Advances in optics, digital computers, digitizers and image processing software, and low-light-level photodetectors have improved the application of FRET microscopy [10], [11], [12]. The development of mutant forms of green fluorescent proteins (GFP), which can be used to reveal the chemical and molecular dynamics of proteins in intact cells, has significantly broadened the usefulness of FRET microscopy, allowing localization and measurement of protein activity [13], [14]. Visualizing fluorescently tagged proteins in living (or fixed) cells with conventional fluorescence microscopy is limited by the contribution of out-of-focus signal from above and below the focal plane [8]. The most widely used techniques to remedy these problems and produce more reliable three-dimensional data are digital deconvolution and confocal microscopy [15], [16]. Laser scanning confocal microscopy (LSCM) uses a pinhole aperture to restrict the out-of-focus flare reaching a detector, the photomultiplier tube (PMT). Increased excitation intensity increases the signal, but also increases photobleaching and photodamage.

These limitations can be overcome with two-photon excitation FRET microscopy (2p-FRET). Two-photon excitation uses a diffraction-limited spot illumination of the specimen, which reduces photobleaching and photodamage outside the area of excitation [17], [18], [19]. Because excitation occurs only at the focal plane, a pinhole is not required and many more photons reach the PMT as compared with confocal microscopy. The use of infrared laser light excitation instead of ultraviolet laser light also reduces phototoxicity in the living cell. The extent of photobleaching with 2p excitation may be greater than one-photon (1p) excitation at the focal plane [20], but overall photobleaching of the sample is considerably reduced in 2p [18]. Further, it is possible to minimize the photobleaching in 2p-FRET microscopy by reducing the average power of the excitation IR laser at the specimen plane.

One of the important conditions for FRET to occur is the overlap of the emission spectrum of the donor with the absorption spectrum of the acceptor [21], [22], [23]. As a result of spectral overlap, the FRET signal is always contaminated by donor emission into the acceptor channel and by the excitation of acceptor molecules by the donor excitation wavelength (see Fig. 1). Both of these signals are termed spectral bleed-through (SBT) signal into the acceptor channel. In principle, the SBT signal is the same for 1p- and 2p-FRET microscopy. In addition to SBT, the FRET signals in the acceptor channel also require correction for spectral sensitivity variations in donor and acceptor channels, autofluorescence, and detector and optical noise, which contaminate the FRET signal.

In this article, we describe the development of 1p- and 2p-FRET microscopy techniques and a methodology to characterize the systems for protein localization. We have described an algorithm for the FRET system that corrects for donor and acceptor SBT and variation in signal intensity due to fluorophore expression (or concentration) levels in living or fixed cells. The energy transfer efficiency (E) and distance (r) between donor and acceptor molecules were calculated (or estimated) using the same cellular image used for FRET imaging.

Section snippets

Polarized epithelial cells

Polarized epithelial MDCK cells, stably transfected with polymeric IgA receptor (pIgA-R), were grown for 3 days in Transwell-Clear inserts, washed with phosphate-buffered saline (PBS), and followed by internalization of 160μg/ml pIgA-R-IgG ligands ([Fab]2 pseudo ligands) conjugated to Alexa 488 (www.probes.com) or Cy3 for 4 h at 17 °C. The ligands were applied to the apical and basolateral plasma membrane (PM), respectively. At 17 °C the pIgA-R-ligand complexes moved into the subapical region [24]

Algorithm

There are various methods to assess the SBT contamination in FRET image acquisition. Donor bleed-through can be calculated and corrected using the percentage of the spectral area of the donor emission spill over into the acceptor emission spectrum or FRET channel (see Fig. 1). In the case of acceptor bleed-through it is difficult to determine the fraction of excitation of the acceptor by the donor wavelength and its emission in the acceptor channel. Moreover, the available methods for FRET data

Comparison of conventional and PFRET methods in the determination of energy transfer efficiency due to variation in fluorophore expression levels (FELs)

It has been demonstrated in the literature that the concentration or uneven labeling of the fluorophore molecule is a common problem in biological signal measurement using fluorescence microscopy techniques [12]. As demonstrated in the literature, ratio imaging (IA/ID) would help to verify whether FRET occurred [8], but it does not provide any information about the energy transfer efficiency or the distance calculation between donor and acceptor molecules. The correction of the unevenness is

Conclusion

In this article we have described the development of confocal (1p-FRET) and two-photon excitation (2p-FRET) FRET imaging microscopy systems. We analyzed the problems associated with FRET microscopy techniques in monitoring and quantifying FRET signals. Our approach to SBT correction is very effective and provides significant statistical data to substantiate the evidence for fixed (MDCK) and live (GHFT1-5) cells. The capability of this algorithm for precision FRET (PFRET) data analysis was

Acknowledgements

We thank Colten Noakes and Cindy Booker for their expert assistance. This work was supported by the W.M. Keck Foundation. We acknowledge Dr. James Demas, Department of Chemistry, and Dr. Martin Straume, Center for Biomathematical Technology, for their helpful discussions.

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