Both the test tube and SDS-gel are powerful platforms for the study of protein-protein interaction but, ultimately, functional relationships between protein binding partners must be understood within the living cell. The advent of fluorescent protein (FP) color mutants derived from the jellyfish Aequorea victoria GFP has made Förster Resonance Energy Transfer (FRET) microscopy a practical tool for the study of protein interaction in a live cell. FRET is a non-radiative process that occurs when the energy of an excited donor fluorophore is transferred to an acceptor fluorophore by dipole-dipole coupling (Förster 1949); in this process, donor fluorescence is quenched and acceptor emission is increased or “sensitized.” For energy transfer to occur, both the donor and acceptor fluorophore must be in close proximity. The efficiency of energy transfer inversely correlates with the distance between fluorophores, increasing as the gap between fluorophores decreases, and can be used to calculate the fluorophore distance, being most accurately measured from 2 to 8 nm. Thus, when an appropriate donor and acceptor FRET pair is used to respectively tag target molecules of interest, both temporal and spatial information can be quantitatively measured below the limit of resolution on a light microscope by virtue of the efficiency of energy transfer from the donor to the acceptor fluorophore. Based on the physics that define FRET, it is natural to assume that FRET is a definitive indicator of protein-protein interaction; yet, it is extremely important to recognize that FRET measurements are rulings of proxy for the interaction of the tagged molecules and, accordingly, in vitro studies remain a critical component for any rationale that drives a FRET-based experiment. But when protein associations are grounded in a supporting body of experimental evidence, FRET becomes a powerful ruler for the study of molecular interaction within a living cell.
Thus far, the preponderance of empirical evidence indicates that the FPs, when expressed on their own or when tagged to a protein of interest, do not induce measurable toxic effects on cells and, quite remarkably, rarely interfere with protein function. While FPs have been placed on either the amino- or carboxy-termini of innumerable proteins, it cannot be overstated that an assessment of tagged-protein function relative to the wild-type protein must be measured experimentally, with the best barometer being rescue of the knockout phenotype. Also, full advantage should be taken of any available structural information to intelligently design the tagged protein: it is of no use to tag an amino-terminus if it is known to be buried within the protein structure or interact with another protein especially when placement of the FP within a flexible loop can be a successful alternative (Janetopoulos, Jin et al. 2001).
In pioneering live-cell FRET experiments, the cyan (CFP) and yellow fluorescent protein (YFP) variants were used to create biosensors to monitor rapid changes in concentration of second messengers such as calcium and cAMP in response to stimuli (REFs). Here CFP and YFP were linked between a specific polypeptide sequence that would change conformation and, thus, the distance between the FRET pair upon binding to the second messenger. While keeping the ratio of the FRET pair fixed at 1:1 greatly simplifies FRET measurements, FRET has been used successfully for the study of transient protein-protein interaction in signal transduction cascades in living cells by tagging individual interacting proteins with either a donor or acceptor FP (Tolar, Sohn et al. 2005; Xu, Meier-Schellersheim et al. 2005; Sohn, Tolar et al. 2006). When well controlled, protein complex formation and dissociation between tagged proteins of interest can be studied at the subcellular level (Schaufele et al., and Wallrabe and Barroso in (Periasamy and Day 2005).
This Unit describes the acceptor-sensitized emission FRET method using a confocal microscope for image acquisition and is intended to complement the acceptor photobleaching FRET method previously described in Unit 19.5 (Wouters and Bastiaens 2001); although, an example of acceptor photobleaching is provided to demonstrate the consistency between the methods on the samples presented below. In contrast to acceptor photobleaching, which is an end-point assay that destroys the acceptor fluorophore, the sensitized emission method is amenable for FRET measurements in live cells and can be used to measure changes in FRET efficiency over time. The purpose of the Unit is to provide a basic starting point for understanding and performing the sensitized emission method with a simple teaching tool for live cell imaging. References that discuss the vagaries of acquiring and analyzing FRET between individually tagged molecules are provided.
Familiarity with DNA transfection techniques, cell culturing and molecular cloning protocols are assumed for this Unit, as is a general understanding of epi-fluorescence microscopy. The authors suggest several excellent web primers (micro.magnet.fsu.edu/; www.olympusmicro.com/; www.microscopyu.com/) for imaging techniques and principles. A step-by-step protocol is provided for cell culturing, and image acquisition using live cells using the Zeiss LSM 510, a laser-scanning microscope equipped with two photomultiplier tubes (PMTs) for detecting emission through selected bandpass filters. Data analysis was performed using the FRET analysis software package provided by Zeiss. While the chapter will be written in a generalized manner referring to “Donor” and “Acceptor” fluorophores, specific examples will be provided using newly developed FRET calibration tools composed of a CFP and YFP variant (Cerulean and Venus respectively) coupled with different lengths of amino acids (Koushik, Chen et al. 2006); these constructs are readily available by written request from the original authors. In addition, the Cerulean-Venus fusion constructs can serve as excellent teaching tools for laboratories and imaging facilities that have FRET-neophytes interested in applying the technique toward their research questions.