Protein–protein interactions are critical to most biological processes, extending from the formation of cellular macromolecular structures and enzymatic complexes to the regulation of signal transduction pathways. The cellular machinery is completely dependent on the exchange of signals, ions, and nutrients between the extracellular environment and the interior of cells. Preexisting and ligand-induced associations of membrane proteins are crucial to such processes. Although generally applied biochemical and immunological approaches (i.e., cocapping, coprecipitation, chemical cross-linking) have provided valuable information about the topography of cell surface protein interactions, such techniques have several disadvantages. For example, the necessary application of extraction or isolation procedures prevent the investigation of proteins in their natural environment and may, on the one hand, disrupt protein–protein interactions or, on the other hand, induce the formation of artificial protein aggregates, the latter being a main concern in case of high expression of cell surface glycoproteins (1). In contrast, fluorescence resonance energy transfer (FRET) offers a suitable and convenient alternative because it can be performed on live cells without major interference with the physiologic condition of the cells. FRET is a process by which an excited donor dye transfers its energy to a nearby acceptor dye through radiationless dipole–dipole interaction (2). The energy transfer efficiency, which is a function of the inverse sixth power of the distance between the donor and acceptor, changes steeply in the range of 1–10 nm and can be used to assess molecular associations (2).
Flow cytometric FRET (FCET) statistically can provide very accurate information on the cell surface distribution of membrane proteins and conformational changes of biologically active molecules. This technique has been applied successfully to a wide range of biological systems, such as monitoring the association state of membrane proteins in immunologically competent cells and various tumor cells; for review, see Szöllősi et al. (3). However, FCET cannot be used easily in cellular systems that show autofluorescence (3). Although FCET does provide information on the lateral organization of molecules with high sensitivity, the technique is not widely applied to cellular systems for two main reasons. First, advanced dual-laser flow cytometric instruments with excitation wavelengths specific for the donor–acceptor pair are required. Although the measurement of FRET-induced donor quenching does not need dual-laser instruments or complicated evaluation, quenching cannot be used for cell-by-cell data analysis of FRET efficiency (3). Second, accuracy and reproducibility of FCET measurements are compromised if the ligands for the fluorescently labeled probes are expressed at low levels. In such cases, the contribution of autofluorescence may be significant relative to the specific signal.
Several reports have offered solutions to diminish the interference of high cellular autofluorescence with the detection efficiency of the specific signal. The high autofluorescence of alveolar macrophages excited with blue light has been quenched by crystal violet (4). Crystal violet, however, may weaken the specific signal. Furthermore, sodium tetrahydridoborate has been used successfully to reduce the autofluorescence of immunochemically stained cells in suspension (5). Alternatively, time-resolved detection of the fluorescence of europium chelates can improve the signal-to-noise (S:N) ratio even in glutaraldehyde fixed samples, where autofluorescence is always high. This is achieved by selective detection of the long-lived europium fluorescence through a time window in which short-lived autofluorescence has already decayed (6). In addition, a mathematical model based on the possible correlation between the autofluorescence and probe fluorescence has been developed for autofluorescence correction in flow cytometry (7). This model, however, is not based on a cell-by-cell approach, and it corrects only the distribution histograms to obtain a better discrimination between the so-called positive and negative cells (7). Importantly, multicolor compensation of autofluorescence on a cell-by-cell basis has been developed for flow cytometric analysis with the use of single-laser (8–11) and dual-laser (12, 13) excitations. These procedures are based on the observation that the excitation and emission spectra of autofluorescence and immunofluorescence are different. In fact, cellular autofluorescence is higher in the blue and green spectral regions than in the red spectrum, so excitation and detection at longer wavelengths reduce the contribution of autofluorescence (14, 15). Newly developed fluorophores with excitation and emission spectra in the red region are therefore superior to the more popular fluorescein and rhodamine fluorophores in FRET experiments. The carbocyanine dyes 3, 5, and 7 (Cy3, Cy5, and Cy7) are good candidates in this respect, and their application has been successful in microscopic FRET measurements (16–19).
We report the development of a highly sensitive FCET method that takes advantage of long emission wavelengths of cyanine dyes and a new technique for cell-by-cell correction of autofluorescence. This new FCET method can be applied in cellular systems where low S:N ratio limits the application of the conventional technique (20). We found that the spectral characteristics of Cy3 and Cy5 as a FRET dye pair sufficiently conform to the optical parameters of the dual-laser FACSCalibur flow cytometer. The dual-laser FACSCalibur is used commonly in daily clinical routine and scientific research, thus facilitating the implementation of the FCET technology. The use of the long emission wavelengths of cyanine dyes markedly reduces the cellular autofluorescence, which in combination with a cell-by-cell correction of autofluorescence using the broad spectrum of autofluorescence, significantly increases the accuracy of energy transfer data analysis.
To date, FRET results obtained by different researchers using distinct labeling protocols (21) are hard to compare. The increased sensitivity of our modified FCET technique allowed us to address important issues, namely the applicability of the method in cellular systems with high autofluorescence and the effect of different direct and indirect immunolabeling approaches on the measured FRET efficiencies. We could detect homoassociation of CD45 isoforms in cells having low expression levels comparable with cellular autofluorescence. We also examined the effects of direct and indirect immunofluorescent labeling with whole immunoglobulin G (IgG) or antigen-binding fragments (Fab) on the efficiency of an intramolecular energy transfer process occurring between the heavy and the light chains of HLA class I molecules. The highest FRET values were measured with directly labeled donor and acceptor antibodies. Increasing the complexity of the labeling scheme by introducing the secondary antibody clearly showed a decrease in the energy transfer efficiency.
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- MATERIALS AND METHODS
- LITERATURE CITED
Although FCET is often used to study molecular associations in intact cells (3, 16, 19), widespread application of the technique is limited by the need for a relatively high expression level of proteins to achieve a high S:N ratio. Cellular autofluorescence is one major source of noise in flow cytometry, thereby decreasing the S:N ratio. In this study we used two approaches to decrease the relative contribution of autofluorescence to the measured intensities: (a) application of long wavelength dyes, whose excitation and emission spectra show less overlap with those of cellular autofluorescence; and (b) a mathematical cell-by-cell correction for autofluorescence with the use of an independent fluorescence channel as a measure for cellular autofluorescence. We demonstrated that the lowest number of receptors, which can be measured reliably by FCET, is significantly lower if the above approaches are applied. In addition, these approaches can be implemented on commercially available instruments. Because secondary labeling strategies are often used to increase the signal in flow cytometry, we carried out a detailed analysis as to whether different labeling protocols change FCET measurements. Our results emphasized that FCET results should be compared with care, when obtained with different labeling methods.
We used Cy3 and Cy5 as the donor–acceptor pair for FCET measurements. Autofluorescence is less of a problem in the red region of the spectrum, which makes autofluorescence almost negligible in the case of Cy5. This makes the Cy3–Cy5 pair superior to the more widely used fluorescein–rhodamine pair for FRET applications. The extent of spectral overlap between Cy3 emission and Cy5 absorption is similar to that between fluorescein emission and tetramethylrhodamine absorption, resulting in similar R0 values (5.0 and 5.6 nm) (26, 27). Even if fluorescein-like dyes with higher fluorescence quantum efficiency and less photobleaching are available, the relatively high autofluorescence of cells is a problem in this spectral region, especially when less abundantly expressed proteins are studied.
Cellular autofluorescence originates mainly from the cytoplasm. Oxidative processes and certain molecules (e.g., reduced nicotinamide adenine dinucleotide (NADH) and catecholamines) may contribute to autofluorescence (28). Therefore, the autofluorescence level depends on the metabolic state of the cell and can display wide heterogeneity even among cultured cells derived from a single progenitor.
Because of the cell-by-cell variation in autofluorescence, methods using a population average value are not accurate to correct for autofluorescence. On the one hand, cells with a lower than average autofluorescence are overcorrected by this method, which, in the case of low specific signal, may even result in negative values. On the other hand, highly autofluorescent cells are undercorrected by this method. Inaccurate correction of fluorescence intensities significantly affects the calculated energy transfer values because the measured parameters are divided by each other, as is necessary in FCET calculations, which results in compounded errors in the derived parameters.
To improve the accuracy of FCET measurements, we used a cell-by-cell correction for autofluorescence. The autofluorescence values measured in the different channels correlated extremely well, which was a prerequisite for the correction on a single-cell basis (Fig. 1). Strongly positive correlations of autofluorescence signals were detected on other T-cell lines (Jurkat) and confluent breast carcinoma cells (SKBR-3, MDA-453; data not shown). Cell-by-cell correction for autofluorescence was especially important for the FL2 and FL3 channels because autofluorescence is not negligible in these channels as it is in the FL4 channel. In addition, miscalculation of the FL2 signal is the largest source of error in energy transfer calculations (29). Improved data analysis resulted in smaller errors in the calculation of energy transfer values of single cells. This contention is supported by the observation that FCET histograms showed a smaller coefficient of variation (Fig. 2), which made the calculation of the mean of FCET histograms more reliable. Reduction of autofluorescence-related errors increases not only the reproducibility of FCET measurements but also the accuracy of the method; i.e., FCET histograms not only get narrower but also may be shifted. This is crucial to the study of proteins with very low expression levels, such as the CD45 protein tyrosine phosphatase on a transfected HPB-ALL T-cell line (Fig. 2). Moreover, the significantly increased sensitivity of our new FCET method incorporating cell-by-cell correction of autofluorescence allowed the detection of small subpopulations with different energy transfer values (Fig. 3).
In many cases, the aim of FRET experiments is to decide whether the proteins of interest associate with each other, i.e., whether the measured FRET efficiency is significantly different from zero. FRET values above 5% are considered significantly different from zero in the case of the fluorescein–rhodamine donor–acceptor pair (25). Smaller coefficients of variation of FCET histograms and increased reproducibility and accuracy of the modified method make lower energy transfer values acceptable as significantly different from zero. Of course, this limit still depends on the S:N ratio, but in optimal conditions it can be as low as 2% (Pascal Batard, personal communication).
Indirect immunofluorescent labeling strategies may be applied to FCET measurements, if a suitable fluorophore-conjugated monoclonal antibody is not available or to enhance the specific fluorescence signal (21). Our intramolecular FCET measurements between the heavy and light chains of HLA class I molecules showed that the application of a larger antibody complex causes a decrease in FRET efficiency due to the geometry of the antibody complexes; i.e., when antibody or Fab complexes get larger, the actual distance between the donor and acceptor fluorophores increases. This explains the decreased FRET efficiency when fluorescent secondary Fabs are used on the donor and acceptor sides relative to primary fluorescent antibodies. It also explains the additional decrease in FRET efficiency when whole fluorescent secondary antibodies were used. The size of the applied fluorescent dyes also can affect the value of energy transfer. Energy transfer efficiencies are even lower when using phycoerythrin- and allophycocyanin-conjugated mAbs (Pascal Batard, personal communication) in comparison with Cy3- and Cy5-conjugated mAbs. Pfeiffer et al. (30) also obtained lower FRET efficiencies when applying phycoerythrin and Cy5 dyes as donor–acceptor pairs. Labeling with fluorescent secondary antibody on the acceptor side not only increases the donor–acceptor distance but also may increase the acceptor concentration because, instead of a single acceptor-labeled primary antibody, several acceptor-labeled secondary antibodies could be present. To rule out this possibility, the secondary antibodies and Fabs were applied in such a concentration that the donor and acceptor concentrations were kept constant. We applied this approach for antibody combinations where we were able to make sure that all epitopes were labeled with at least one fluorescencently tagged IgG or Fab to avoid the disturbing effect of empty epitopes in the FRET efficiency calculation. In these cases, the FRET values are not influenced by the donor–acceptor ratio but by the relative orientation of the fluorophores and the distance between them. Secondary labeling with fluorescent Fab or IgG on the donor side decreases FRET efficiency because the size of the labeling complex increases (Table 1). Although it is difficult to predict the value of energy transfer without molecular models and calculations, our results emphasized that FRET values cannot be directly compared with each other if they were obtained with different labeling strategies.
In conclusion, we provided a detailed description of a modified method to measure energy transfer in flow cytometry. The main value of the approach is reduction of autofluorescence-related errors, which is achieved by the application of long wavelength dyes, such as Cy3 and Cy5, as a donor–acceptor pair and by cell-by-cell correction of autofluorescence. Our modified FCET method allows energy transfer efficiency to be determined on cellular systems with a very low expression level of the molecules. Further, the lower variance of FCET distributions enables a much more accurate discrimination of subpopulations having distinct FRET efficiencies. The practical advantage of the new method is that it can be implemented easily on a commercially available dual-laser benchtop flow cytometer without the need for hardware modifications.