Simultaneous FRAP, FLIM and FAIM for measurements of protein mobility and interaction in living cells

: We present a novel integrated multimodal fluorescence microscopy technique for simultaneous fluorescence recovery after photobleaching (FRAP), fluorescence lifetime imaging (FLIM) and fluorescence anisotropy imaging (FAIM). This approach captures a series of polarization-resolved fluorescence lifetime images during a FRAP recovery, maximizing the information available from a limited photon budget. We have applied this method to analyse the behaviour of GFP-labelled coxsackievirus and adenovirus receptor (CAR) in living human epithelial cells. Our data reveal that CAR exists in oligomeric states throughout the cell, and that these complexes occur in conjunction with high immobile fractions of the receptor at cell-cell junctions. These findings shed light on previously unknown molecular associations between CAR receptors in intact cells and demonstrate the power of combined FRAP, FLIM and FAIM microscopy as a robust method to analyse complex multi-component dynamics in living cells. the FRAP experiment. During our measured fluorescence recovery unbleached CAR-GFP diffuses into the ROI. We attribute the overall decrease in the anisotropy as a function of time during the recovery to an increase in the proportion of GFP units undergoing FRET, with dimeric/oligomeric units diffusing into the ROI. We note also


Introduction
Fluorescence microscopy is an essential tool for live cell imaging yielding information on macromolecular structure, location, interactions and dynamics. The fluorescence spectrum, intensity, lifetime, polarization and their evolution in time can provide a wealth of information about intracellular environments and their dynamics [1][2][3]. Capturing as many of these features as possible in a single measurement maximises the information extracted from the sample and ensures that the limited photon budget available from each fluorophore is used effectively. Living cells show dynamic behaviour over a wide range of timescales -from subnanosecond conformational fluctuations to chemical reactions and interactions over several hours [4]. Protein mobility can be measured using a variety of fluorescence microscopy techniques. For example, fluorescence recovery after photobleaching (FRAP), in which the fluorescence intensity in a region of interest is measured as a function of time following rapid, irreversible photobleaching, can report on translational intracellular protein mobility [5,6]. Such experiments are relatively straightforward to perform and methods for analysis are becoming increasingly more sophisticated, with general models for calculations of diffusion coefficients from confocal FRAP experiments [7][8][9]. The rotational mobility of fluorophores can be probed by measuring the emission polarized parallel and perpendicular to the excitation light and calculating the fluorescence anisotropy. This approach can also be used to identify Förster resonance energy transfer (FRET) between both different fluorophore types [10][11][12] and between the same type of fluorophores (i.e. homo-FRET or emFRET) [13][14][15][16][17][18][19][20][21][22][23]. In fact, with rare exceptions, e.g. [24], fluorescence anisotropy can be the only method to identify homo-FRET if the two fluorophores are in the same environment and their lifetimes are identical. Time-resolved fluorescence anisotropy imaging (tr-FAIM) [17,[25][26][27] is currently rarely used in biology but has potential to be a powerful indicator of rotational diffusion of fluorophores, protein structure or function [1]. Underpinning tr-FAIM, fluorescence lifetime imaging microscopy (FLIM) [2,3,[28][29][30][31][32][33] maps the fluorescence lifetime in every pixel of an image and is a powerful technique for probing the local environment of a fluorophore as the measured lifetime is largely independent of fluorophore concentration, but can be sensitive to pH [34], refractive index [35][36][37][38], reactive quenching species [39] and viscosity [30,31,[40][41][42]. Both fluorescence anisotropy and FRAP have previously been used independently for a study of aggregation states of alpha-synuclein, a protein which plays a role in Parkinson's disease [43], and an arrangement for dynamic FRAP and rotational diffusion measurements for colloids has been presented [44]. FRAP and FLIM have also recently been used independently in a study of keratinocyte migration [45], influenza virus association with lipid rafts [46], and cyanobacteria [47]. FLIM and anisotropy have recently been used for high content screening [48], and a system for widefield spectrallyresolved fluorescence lifetime and anisotropy imaging has been presented [49]. However, to our knowledge there has not yet been an example of FLIM, FRAP and FAIM used simultaneously in combination for studies of protein dynamics.
In living samples undergoing continuous change it is desirable to extract the maximum amount of information available whilst minimising exposure to intense light and reducing the effects of photobleaching, phototoxicity and varying sample morphology due to motion on the measurements by keeping acquisition times short. As such, it is beneficial to combine measurements rather than perform many consecutive measurements on a sample. To address this issue and in an effort to maximise the amount of information obtained from a single experiment we have combined three fluorescence microscopy techniques [50], as schematically illustrated in Fig. 1. Our experimental set-up is based on an inverted laser scanning confocal microscope with time-correlated single photon counting (TCSPC) detection using hybrid GaAsP photomultiplier detectors. This allows for FLIM, FRAP and tr-FAIM experiments to be performed simultaneously, rather than sequentially, for the same photon budget, and with the same acquisition times and spatial resolution as a typical FLIM experiment, whilst retaining all of the benefits of each individual fluorescence technique. The time-resolution for the FRAP recovery curves is on the timescale of seconds which is well suited to monitoring protein dynamics in membranes. The anisotropy measurements allow for identification of species undergoing FRET and the fluorescence lifetime measurements have the potential to allow identification of local variations in the fluorophore environment. This approach is useful, for example, for distinguishing between high local fluorophore concentrations and oligomeric structures. We have measured fluorescence lifetimes, depolarization, and translational mobility of coxsackievirus and adenovirus receptor (CAR) [51] labelled with GFP in living human bronchial epithelial cells (HBEC) simultaneously in the same experiment. CAR is a transmembrane protein that localises to cell-cell junctions and is proposed to play a role in mediating cell-cell adhesion [52,53]. The extracellular N-terminal domain of CAR is known to exist as a homo-dimer in solution and various biochemical evidence suggest this may contribute to cis and trans interactions between receptors, i.e. within the same cell and across cell-cell junctions [51][52][53][54]. However, the receptor state in intact cells and the potential role of self-association in controlling cell-cell adhesion and adenovirus docking is currently unknown [54,55]. We have therefore applied combined FRAP, FLIM, tr-FAIM microscopy to investigate the dynamics and dimerisation of CAR in living cells.

Preparation of rhodamine 123 samples
All materials were used as received and solvents were spectrophotometric grade. A stock solution of 1.3 mM rhodamine 123 (rh123, Mw = 380.82, Sigma, UK) in methanol (Sigma Aldrich, UK) was made and 40 μl of the stock solution was added to a 10 ml mixture of glycerol (Sigma Aldrich, UK) and methanol with volume fraction 90:10 to give a final concentration of the dye 5.2 μM. For imaging, 200 μl of the solution was imaged in one well of a 96-well plate with a coverglass underside (Whatman) at room temperature.

Cell culture and preparation
Cells were cultured on a 6-well plate as part of a resistively-heated micro-incubation system (SmartSlide50, Wafergen, UK). For imaging the cells were heated to 37 °C and 5% CO 2 / 95% air was flowed through the well. Immortalised human bronchial epithelial cells (HBEC) were a gift from Dr Jerry Shay (UT Southwestern [56]) and were grown in keratinocyte serum-free media (KSFM; Invitrogen). CAR-GFP expressing stable cell lines were produced using lentiviral expression. CAR-GFP lentivirus particles were generated in 293T packaging cells (as in ref [53].) and these cells were maintained in DMEM containing 10% FCS, supplemented with glutamine. Plasmids encoding full-length CAR have been described previously [57]. Full length CAR-GFP was cloned in frame into pHR9SIN-SEW lentiviral expression vector, which was a gift from Dr Adrian Thrasher (Institute of Child Health, UCL, London [58]), and into pGEX-2T. Cells were plated at high density onto custom designed 6well plates (SmartSlide50, Wafergen, UK) 36 hours prior to analysis. For control experiments HBEC were transiently transfected with eGFP-N1 (Clontech) using Fugene 6 (Roche) according to the manufacturer's instructions and imaged 36 hours post-transfection.

Combined FRAP, FLIM, tr-FAIM microscopy
The microscopy experiments were performed using an inverted confocal laser scanning microscope (Leica TCS SP2). Samples were imaged using a 63 × water immersion objective (NA 1.2, heated to 37 °C) with a line scan speed of 400 Hz (1.64 s per frame) Two lasers were used for the FRAP experiment -a pulsed diode laser at 467 nm (Hamamatsu PLP 10) with pulse duration of 90 ps, repetition rate of 20 MHz and average power ~1μW for the preand post-bleach imaging, and a continuous wave Ar + laser at 488 nm with an average power of ~1 mW for the bleach frame. A time-lapse acquisition series was set up with three prebleach frames, followed by one bleach frame of duration 1.64 s, and then post-bleach frames which were looped until the FRAP recovery was complete and the image acquisition was terminated. The repetition rate of the diode laser gave a 50 ns window for acquisition of fluorescence decays and thus the advantage of being able to record complete decays from rh123 and GFP. The fluorescence was passed through a polarizing beamsplitter cube and the orthogonally polarized components were detected using two GaAsP hybrid detectors measurements of HBEC expressing CAR-GFP and control measurements of HBEC expressing empty vector GFP constructs were performed with an upgraded system using two SPC-150 photon counting boards (Becker & Hickl).

Data analysis
Analysis of the data was performed using TRI2 time-resolved image analysis software [59]. For the FRAP recovery curve each pair of images recorded in each frame was summed to give an image of the total measured intensity per frame. The bleached ROI was masked and the intensity was obtained by adding the photon counts. The intensity was corrected for bleaching and fluctuations by measuring the intensity in another region of the image where the cells were immobile on the time scale of the experiment. The recovery curves were fitted to a monoexponential recovery curve. The FLIM images were created by summing all of the measured parallel and perpendicular frames, respectively, and combining them according to the denominator of Eq. (1). We have verified that even for a high NA microscope objective this is valid for fluorescence lifetime measurements. A 50 ns window for fluorescence decays ensures that we measure complete fluorescence decays from GFP. Steady-state anisotropy images were created using the summed parallel and summed perpendicular images an anisotropy macro in TRI2 which processed the images according to Eq. (1). Time-resolved anisotropy decays were measured by integrating the fluorescence over all pixels in the time and polarization images. For FLIM binning of 7x7 pixels was used.
A FRAP recovery curve is measured from a region of interest (ROI) using intensity values from pairs of time and polarization-resolved fluorescence images recorded at regular intervals following photobleaching. Fluorescence anisotropy maps are produced by processing the summed images for each measured polarization component according to Eq. (1), where Ipara and I perp are the intensity values for fluorescence polarized parallel and perpendicular to the incident radiation, respectively, and G accounts for the difference in detection efficiency for the two measured polarization states.
Combining the time and polarization-resolved images according to the denominator of Eq. (1) and fitting the fluorescence decays in each pixel using an exponential decay model yields a FLIM image. Therefore, during the course of the polarization-resolved FLIM measurements it is possible to extract quantitative translational and homo-FRET oligomerization data and measure fluorescence lifetimes and homo-FRET without extending the acquisition or exposure times used for a typical confocal FLIM experiment.

Generation of rotational correlation time image and histogram
The rotational correlation time image and histogram for the homogeneous solution sample of rh123 were generated using OptiSP e C software from Becker & Hickl. Briefly, the software is capable of summing multiple time-resolved image files, and we used this function to import and sum 150 images from each of the parallel and perpendicular polarization detection channels. Using spatial binning of 15x15 pixels and a G-factor of 1.35 the fluorescence decays recorded for each polarization in each pixel were fitted simultaneously according using Eq. (2) and Eq. (3) [61].
[ ] where || F and F ⊥ are the fluorescence decays for polarizations parallel and perpendicular to the incident radiation, respectively. I(t) and r(t) are the time-dependent fluorescence intensity and anisotropy, respectively, G accounts for the difference in detection efficiencies of each polarization component, and R is the instrument response function (IRF) with t s the time shift between the IRF and the fluorescence data.

Combined FRAP, FLIM, tr-FAIM microscopy of rhodamine 123 in solution
We first used the combined microscopy set-up to measure fluorescence lifetime, τ f , and diffusion properties of the well-characterised dye rh123 in a 90:10 (v/v) mixture of glycerol and methanol (viscosity, η = 400 ± 30 cP, refractive index, n = 1.46). The FLIM image ( Fig.  2(a)) shows a uniform distribution of fluorescence lifetimes as expected from a dye in homogeneous solution. A typical rh123 fluorescence decay, with τ f , = 3.19 ± 0.07 ns (χ 2 = 1.15), is shown in Fig. 2(a) (inset). The fluorescence lifetime histogram is symmetric and well fitted by a Gaussian distribution centred around 3.25 ns with a full width at half maximum (FWHM) of 0.4 ns. This is in good agreement with the expected value in a high refractive index methanol/glycerol medium (n = 1.46) based on the Strickler-Berg relationship between lifetime and refractive index and the reported lifetime in ref [60]. The steady-state fluorescence anisotropy, r, image is shown in Fig. 2(b). This was created by summing the series of images recorded for each polarization state (parallel and perpendicular) and calculating the anisotropy from the intensity values, || I and I ⊥ , in each pixel using Eq. (1). The image is similarly uniform and the fluorescence anisotropy histogram (Fig. 2(b)) is also symmetric and can be fitted to a Gaussian function centred around 0.236 with a FWHM of 0.07. A map of the rotational correlation time (average tumbling time), τ c , is created for the solution sample (Fig. 2c) by fitting of the polarization-resolved fluorescence decays using OptiSP e C software (Becker & Hickl,) with pixel binning of 15 x 15 pixels. This software allows the determination of rotational correlation times for freely rotating fluorophores by simultaneously fitting both of the polarization-resolved fluorescence decays (Eq. (2) and Eq.
(3)) in each pixel [61]. The image shows a homogeneous distribution of rotational correlation times, τ c , with a peak in the histogram at ~15 ns. The value of τ c for the dye in the solution, obtained from a monoexponential fit to the fluorescence anisotropy decay curve (Fig. 2(e)) with integrated pixel intensities over the whole image is τ c = 15.2 ± 2.8 ns. Using Eq. (4) where k b is the Boltzmann constant, T is absolute temperature, η is viscosity, and r is the hydrodynamic radius of the dye, the calculated rotational diffusion coefficient, D rot = 0.011 ± 0.002 ns −1 . The initial anisotropy, r 0 = 0.28 ± 0.01 is lower than the theoretical maximum of 0.4 for single photon excitation due to depolarization by the high numerical aperture objective, an effect that has been observed previously [17,62,63]. The FRAP recovery curve ( Fig. 2(d) with selected images from the recovery image series, inset) can be adequately fitted by a single exponential function with a half time for the fluorescence recovery, t 1/2 = 11.5 ± 0.5 s and no immobile fraction, as expected for a dye in solution. This is indicative of free diffusion. Assuming Stokes-Einstein-Debye behaviour and using Eq.
where γ is a beam geometry parameter (γ = 0.88 for a circular beam bleach), w is the radius of the bleach area and t 1/2 is the half time of the fluorescence recovery, the calculated translational diffusion coefficient is D trans = 1.8 ± 0.2 μm 2 s −1 .
The radius, r, of the rh123 molecule was calculated from the rearranged Stokes-Einstein-Debye equations for translational and rotational diffusion (Eq. (6) and Eq. (7)), yielding values of ~3.1 ± 0.4 Å and ~3.4 ± 0.2 Å for the radius of rh123, respectively, which are in good agreement with values calculated from literature values of the diffusion coefficient of rhodamine dyes [64,65].

Combined FRAP, FLIM, tr-FAIM microscopy of CAR-GFP in living human epithelial cells
Having validated our set-up on rh123 in solution, we used it to measure the fluorescence lifetimes, FRAP recovery curves and time-resolved fluorescence anisotropy of CAR-GFP stably expressed in living human bronchial epithelial cells (HBEC). For an acquisition time of ~450 s we recorded a series of 150 pairs of time-and polarization-resolved confocal fluorescence images simultaneously during a single FRAP experiment. The CAR-GFP signal was localised to both cell-cell junctions and within perinuclear compartments in line with previous studies [53,57]. The tagged receptor has also previously been shown to function in an identical fashion to untagged CAR and fully supports adenovirus type5 infection, binding to known interaction partners like multi PDZ domain protein 1 (MUPP-1), and cell-cell junction formation [57,66].  [35,36,67,68] and our previous FLIM measurements of CAR-GFP alone [57]. The steady-state anisotropy image (Fig. 3(d)) shows a narrow distribution of anisotropy values around the peak at 0.215 (histogram, Fig.  3(f)). Importantly, there is also no discernible difference in the anisotropy values between the junctions and the perinuclear regions. As a control we measured HBEC expressing empty vector monomeric GFP and found that the average anisotropy value in the cells, r = 0.308 ± 0.012 compared with r = 0.214 ± 0.048 at cell-cell junctions in HBEC expressing CAR-GFP ( Fig. 3(i)). These values indicate the presence of a population of fluorophores in the cells expressing CAR-GFP undergoing more rapid depolarization than monomeric GFP, and this is attributed to rapid depolarization via non-radiative energy transfer between GFP units, i.e. homo-FRET. By using the parallel and perpendicular fluorescence decays from all pixels in the summed series of the measured orthogonally polarized fluorescence images we constructed the timeresolved anisotropy decay (Fig. 3(g)). The decay can be fitted well using a monoexponential function giving an apparent rotational correlation time of 5.99 ± 1.16 ns, and a limiting anisotropy, r∞ = 0.15 ± 0.01, indicative of homo-FRET between adjacent GFP units. The existence of homo-dimers or higher order oligomers can decrease the measured anisotropy decay time due to energy transfer between adjacent fluorophores. The r∞ is then related to the number of GFPs taking part in homo-FRET. Non-radiative resonance energy transfer between adjacent GFP units occurs more rapidly than the rotation of a dimeric/multimeric units leading to more rapid depolarization of the fluorescence than would be seen by rotation alone -an effect which has been exploited for study of protein structure [14] and cluster sizes [13]. It should also be noted that a monoexponential fit to the data is based on the assumption that the rotational correlation time of the constituent GFP units is much longer than the depolarization time due to homo-FRET. Although our time windows for GFP decay measurements are long enough to measure complete GFP fluorescence decays, little Brownian rotational motion takes places during the excited state lifetime. In this case the value of r∞ is influenced by the proportion of monomeric GFP units in the measured volume. In a system including monomers and oligomers with the oligomers undergoing homo-FRET, the anisotropy increases with an increasing proportion of monomers [69]. The value of r∞ has also been seen to decrease with increasing cluster size and can be used as a measure of cluster size in some cases [13]. Our data is therefore consistent with the hypothesis that CAR exists predominantly in nonmonomeric complexes in living epithelial cells, and that this is not restricted to junctions between adjacent cells.
The representative FRAP recovery curve (Fig. 3(h)) measured from the junction ROI outlined as a red circle in Fig. 3(a) shows an incomplete fluorescence recovery and an immobile fraction of ~85%. This is indicative of CAR that is tethered at the junction either through homo-dimerization in cis or trans, or potentially through interactions with other scaffold proteins. Interestingly, the immobile fraction of CAR measured here is remarkably similar to that of the classical adherens junction receptor E-cadherin [70] suggesting adhesion receptors can exhibit similar dynamics at cell-cell junctions or that these receptors are functionally linked as we have previously suggested [66]. The representative recovery curve for the CAR-GFP can be adequately fitted using a monoexponential recovery function, with a recovery half-time, t1/2 = 145 ± 25 s.
Polarization and time-resolved fluorescence decay data is recorded in every frame of the FRAP recovery, yielding polarization-resolved FRAP recovery curves. The curves measured for the rh123 in solution (Fig. 4(a)) and the tight junction CAR-GFP (Fig. 4(c)) are markedly different due to the faster recovery of the fluorescence in the dye solution and the higher number of photons in the ROI. Using these curves it is possible to measure the anisotropy in the bleached ROI as the fluorescence recovers. As expected, the value of the anisotropy in the ROI for the dye solution is invariant ( Fig. 4(b)) due to the uniform distribution of dye molecules and constant viscosity throughout the sample, while the anisotropy can be seen to decrease as a function of recovery time in the CAR-GFP sample ( Fig. 4(d)). We measure this decrease by summing the fluorescence intensity in the ROI for ten adjacent frames in both parallel and perpendicular polarization detectors, and we calculate the error in the anisotropy measurements based on the number of detected photons using the treatment presented by Lidke et al. [71] The time-dependent steady-state anisotropy decrease can be fitted well to a monoexponential decay function with a decay time equal to that measured for the FRAP recovery curve.
Variations in steady-state anisotropy as a function of the fraction of the fluorophores in the ROI which has been bleached have been presented by other authors [19,69]. For the case of randomly distributed monomeric units in the region of interest the anisotropy should remain constant during photobleaching. For a system undergoing FRET photobleaching of acceptor molecules increases the measured anisotropy. CAR-GFP molecules within the ROI are photobleached during the FRAP experiment. During our measured fluorescence recovery unbleached CAR-GFP diffuses into the ROI. We attribute the overall decrease in the anisotropy as a function of time during the recovery to an increase in the proportion of GFP units undergoing FRET, with dimeric/oligomeric units diffusing into the ROI. We note also that the fluorescence lifetime in the ROI remains invariant during the recovery (Fig. 4(d), inset) which is an indication that there is no fluorescence quenching occurring as the local concentration of unbleached fluorophores increases.

Discussion
Fluorescence microscopy techniques offer a minimally invasive means of extracting steadystate and dynamic properties of intracellular environments. Individually, FRAP measurements can report on translational protein mobility and binding, and time-resolved fluorescence anisotropy measurements can report on rotational mobility and clustering. The translational diffusion coefficient is influenced by the viscosity of the medium, the size of the mobile unit, and crowding of the environment which reduces the mean free path. The fluorescence anisotropy is also influenced by the viscosity of the medium, the size of the rotating unit, and energy transfer. Our experimental arrangement combines these measurements, and additionally measures the fluorescence lifetime in a single, simultaneous measurement. As such, potential ambiguities in the interpretation of independently measured FRAP and anisotropy data are eliminated, and more detailed clustering and binding information can be obtained.
In the case of CAR-GFP, the FRAP recovery reveals information that may otherwise be open to misinterpretation from anisotropy experiments alone. A translationally immobile fraction at the cell-cell junctions due to trans dimerisation would also lead to a reduction in rotational mobility at the junctions in comparison to that in the perinuclear regions where it is highly likely that only cis dimerisation would occur. However, this is not evident in the anisotropy image or histograms and the large volume of the dimer unit would yield an anisotropy decay component that is too long to measure using GFP emission even if it were freely rotating. Simply knowing the steady-state anisotropy value in an image cannot reveal more complex dynamics occurring due to hindered rotations and depolarization due to energy transfer. Time-resolved fluorescence anisotropy measurements are essential to disentangle these phenomena and are therefore an important measurement modality integrated into our microscopy set-up. In the case of homo-dimers the homo-FRET contribution to the anisotropy decay is dominant. Similarly, the presence of dimers cannot be unequivocally determined by FRAP measurements. Translational motion can be slowed by crowded or viscous environments as well as an increase in the size of the mobile unit, and an immobile fraction could also be observed for monomeric structures. Equally, anisotropy measurements alone cannot report on the translational mobility of dimers or oligomerized proteins. This highlights the importance of the FRAP component of the measurements to avoid ambiguities and provide a more complete picture of protein behaviour in living cells. Measurement of the fluorescence lifetime of GFP allows for identification of areas where there are variations in the local environment or self-quenching due to high local concentrations. In the case of our experiments the invariant fluorescence lifetime of GFP during the FRAP recovery is suggestive of there being no concentration-dependent fluorescence quenching. Our combined FRAP, FLIM, tr-FAIM microscopy data show that CAR exists in a nonmonomeric state essentially independent of its location -either in perinuclear vesicles or at cell-cell junctions -with a fraction of the population immobile at the junctions. This implies that dimeric complexes are not formed exclusively at the junctions but rather that CAR exists in a multimeric form everywhere in the cell. This has important implications for understanding how CAR is regulated during epithelial cell junction formation, dissolution and response to adenoviral infection. Interestingly, CAR has also recently been shown to undergo clathrin-mediated endocytosis through binding to AP adaptor proteins [72], and our data would also suggest this may occur through internalisation of a dimeric receptor. Our method, outlined here, is highly suited to the analysis of dimeric/oligomeric complexes and transport properties of other receptors, such as G-protein coupled receptors [73], in living cells, and we believe that it represents a valuable addition to the fluorescence microscopy techniques available to study living cells.

Conclusion
We have demonstrated the combination of three powerful fluorescence microscopy techniques -FRAP, FLIM, FAIM -in a single experiment. Simultaneously measuring multiple fluorescence parameters over time -namely position, intensity, lifetime and polarization -yields more insight than sequential measurements using multiple techniques as we examine not only the same cell but the same fluorescence signal, from the same pixels at the same time. The method thus maximises the information available from a limited fluorescence photon budget. It can measure complementary properties required for a comprehensive interpretation of protein dynamics in living cells, which each method on its own cannot provide. We have shown this by measuring the anisotropy as a function of time during a FRAP recovery. This integrated multimodal fluorescence technique thus allows for identification of protein dimers or oligomers and measurement of their translational mobility around the cell. We have demonstrated the power of the technique for monitoring the clustering and dynamics of GFP-tagged cell-cell adhesion molecule CAR, and reveal the fractional preferential multimeric nature of this receptor in intact living cells. Where oligomerization is evident, via low values for fluorescence anisotropy, it is most likely that CAR is interacting in cis within intracellular compartments, but both in cis and trans at cellcell junctions.