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Imaging Forster Resonance Energy Transfer Measurements of Transmembrane Helix Interactions in Lipid Bilayers on a Solid Support† Edwin Li and Kalina Hristova* Department of Materials Science and Engineering, Johns Hopkins University, Baltimore, Maryland 21218 Received May 28, 2004. In Final Form: July 16, 2004 We utilize supported lipid/protein bilayers to probe the dimerization of transmembrane (TM) helices in a membrane environment. The bilayers are formed by incubating substrates with liposomes containing the proteins, and are characterized using fluorescence recovery after photobleaching and imaging Forster resonance energy transfer (FRET). We show that the FRET signal, as a measure of TM helix dimerization, is the same in suspended liposomes and in surface-supported bilayers. This work is the first step toward the development of a new tool for probing the association of TM helices in lipid bilayers.
Introduction Approximately 20% of the open reading frames in complex organisms encode membrane-associated proteins.1 These proteins play key roles in cell adhesion, recognition, motility, energy production, transport of nutrients and cholesterol, and biochemical signal transduction.2 Membrane proteins are the sensors of the living cell, and their potential in biotechnological applications is enormous.3 Engineering of surface-immobilized bilayers containing fully active integral membrane proteins can lead to the design of a new generation of drug screening devices, mimetics of cell and tissue surfaces, and matrices for stress-free cell immobilization. Recent progress in both membrane protein biophysics and planar bilayer methodology is setting the stage for the development of new tools for biomedical research. Here we utilize supported lipid/protein bilayers to probe transmembrane helix dimerization in a membrane environment. Such measurements of helix dimerization are needed for understanding the physical principles underlying vital cellular processes such as membrane protein folding and signal transduction across the plasma membrane. For instance, receptor tyrosine kinases (RTKs) transduce biochemical signals across the plasma membrane via lateral dimerization.4 RTKs exist in a monomer-dimer equilibrium, with the monomer being inactive, while the dimer is active.5 The right balance of monomers and dimers ensures normal cell signaling, and amino acid mutations that inhibit or enhance dimerization often cause pathologies.6 * To whom correspondence should be addressed. E-mail:
[email protected]. Phone: (410) 516-8939. Fax: (410) 516-5293. † Abbreviations: TM, transmembrane; FRAP, fluorescence recovery after photobleaching; FRET, Forster resonance energy transfer; FGFR3, fibroblast growth factor receptor 3, RTK, receptor tyrosine kinase; GPDES, (3-glycidoxypropyl)dimethylethoxysilane; APDES, (3-aminopropyl)dimethylethoxysilane; POPC, 1-palmitoyl2-oleoyl-sn-glycero-3-phosphocholine; NBD-PE, N-(7-nitrobenz2-oxa-1,3-diazol-4-yl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine; HFIP, hexafluoro-2-propanol; MeOH, methanol. (1) Liu, J. F.; Rost, B. Protein Sci. 2001, 10, 1970-1979. (2) von Heijne, G. J. Mol. Biol. 1999, 293, 367-379. (3) Sackmann, E.; Tanaka, M. Trends Biotechnol. 2000, 18, 58-64. (4) Bormann, B. J.; Engelman, D. M. Annu. Rev. Biophys. Biomol. Struct. 1992, 21, 223-242. (5) Schlessinger, J. Cell 2002, 110, 669-672.
Recently, it has become clear that the TM helices of RTKs play a key role in the dimerization process, as suggested by the finding that a single amino acid substitution in the TM helix can lead to constitutive receptor activation and therefore to pathologies.6,7 For example, multiple single amino acid mutations in the TM helix of one RTK, fibroblast growth factor receptor 3 (FGFR3), cause various cancers and developmental abnormalities by affecting the lateral dimerization in the membrane.8-11 In our laboratory we seek to understand how these FGFR3 mutations alter the thermodynamics of dimerization and cause disease. The first step toward this goal is characterizing the dimerization propensity of the wild-type (normal) FGFR3 TM helix using Forster resonance energy transfer (FRET). In this work, we incorporate the FGFR3 TM helix in surface-supported bilayers and measure the FRET signal as a reporter of helix dimerization. The formation of lipid-only bilayers on a solid support is a well-developed methodology.12-15 Liposomes are incubated with glass substrates, leading to the formation of single bilayers adsorbed on glass, on top of a “cushion” of water molecules. One may expect that the incubation of liposomes containing proteins will lead to the formation of mixed lipid/protein bilayers, such that the transmembrane proteins will be embedded in the substratesupported lipid matrix. However, very little is known about the structure of these bilayers. Are proteinsubstrate and lipid-substrate interactions the same? How (6) Hynes, N. E.; Stern, D. F. Biochim. Biophys. Acta 1994, 1198, 165-184. (7) Miloso, M.; Mazzotti, M.; Vass, W. C.; Beguinot, L. J. Biol. Chem. 1995, 270, 19557-19562. (8) Webster, M. K.; Donoghue, D. J. EMBO J. 1996, 15, 520-527. (9) Vajo, Z.; Francomano, C. A.; Wilkin, D. J. Endocr. Rev. 2000, 21, 23-39. (10) Shiang, R.; Thompson, L. M.; Zhu, Y.-Z.; Church, D. M.; Fielder, T. J.; Bocian, M.; Winokur, S. T.; Wasmuth, J. J. Cell 1994, 78, 335342. (11) van Rhijin, B.; van Tilborg, A.; Lurkin, I.; Bonaventure, J.; de Vries, A.; Thiery, J. P.; van der Kwast, T. H.; Zwarthoff, E.; Radvanyi, F. Eur. J. Hum. Genet. 2002, 10, 819-824. (12) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 25542559. (13) Vontscharner, V.; McConnell, H. M. Biophys. J. 1981, 36, 409419. (14) Sackmann, E. Science 1996, 271, 43-48. (15) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105-113.
10.1021/la048676l CCC: $27.50 © 2004 American Chemical Society Published on Web 09/08/2004
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do protein interactions within the surface-supported bilayer compare to protein interactions within the freely suspended liposomes? As a first step toward characterizing the architecture of surface-supported protein/lipid bilayers, we present fluorescence recovery after photobleaching (FRAP) and FRET measurements in these systems. We show that the TM helices interact strongly with the substrate, and are immobile on both clean and silanized glass substrates. On the other hand, lipid mobility varies and can be controlled, depending on the pretreatment of the glass surfaces. We further show that the FRET signal is the same in suspended liposomes and in surface-supported bilayers. The FRET signal is a measure of protein interactions (in this case dimerization), suggesting that bilayer formation on a solid support does not alter protein interactions. We demonstrate that the measured FRET signals on all studied surfaces are reproducible and identical within experimental error. For our measurements we have utilized and further developed an imaging FRET methodology16-19 that may provide a less expensive and less tedious alternative to solution FRET measurements. Materials and Methods Glass Substrates. Microscope glass slides (Fisher) were cleaned by sonication for 10 min in 2-propanol (IPA), then acetone, and IPA. After the second sonication with IPA, slides were rinsed with deionized water (DI H2O), and immersed in a solution of 70% sulfuric acid (H2SO4) and 30% hydrogen peroxide (H2O2) for 20 min. Slides were then extensively rinsed with DI H2O, and dried under a stream of filtered nitrogen gas. Substrates were silanized overnight in a 10 mM solution of either (3-glycidoxypropyl)dimethylethoxysilane (GPDES) or (3aminopropyl)dimethylethoxysilane (APDES) (Gelest) in xylenes at room temperature. Silanized substrates were rinsed with xylenes, followed by chloroform, and dried under a stream of nitrogen gas. FRET Dyes. A requirement for every donor-acceptor pair of fluorophores for FRET measurements is that the emission spectrum of the donor overlaps with the excitation spectrum of the acceptor. In addition, for imaging FRET the two excitation and the two emission spectra should be well separated, such that the two dyes can be independently excited and observed. The dye pair Cy3 and Cy5 (Amersham) is one of the available pairs suitable for imaging FRET studies. In our experiments Cy3 and Cy5 were excited and observed independently by selecting the appropriate Cy3 or Cy5 filters. Labeling of Proteins. The FGFR3 TM helix, amino acid sequence DEAGSVYAGILSYGVGFFLFILVVAAVTLCRLR, was custom synthesized at the Kansas State Biotechnology Facility. The peptides were purified using reversed-phase HPLC and a water/acetonitrile gradient. The correct molecular weight was confirmed using MALDI-TOF mass spectrometry. The helicity of the proteins was confirmed using CD (see Figure 1B). The single Cys residue in the protein was labeled with either Cy3maleimide or Cy5-maleimide (Amersham). For the labeling reaction, approximately 2 mg of protein was dissolved in 400 µL of hexafluoro-2-propanol (HFIP) and 800 µL of 10 mM phosphate buffer at pH 7. The dyes were first dissolved in 50 µL of methanol (MeOH) and then mixed with the protein. The reaction was carried out at room temperature with frequent mixing for the first 30 min, and then left in the refrigerator overnight. Excess dye was removed using a C2 solid-phase extraction column (16) Kenworthy, A. K.; Edidin, M. J. Cell Biol. 1998, 142, 69-84. (17) Kenworthy, A. K.; Petranova, N.; Edidin, M. Mol. Biol. Cell 2000, 11, 1645-1655. (18) Bastiaens, P. I.; Jovin, T. M. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 8412. (19) Bastiaens, P. I.; Majoul, I. V.; Verveer, P. J.; Soling, H. D.; Jovin, T. M. EMBO J. 1996, 15, 4246-4253.
Li and Hristova (Varian) in 30% acetonitrile. The labeled proteins were eluted in 100% acetonitrile, and further purified by HPLC. Samples collected from the HPLC column (Varian) were lyophilized and redissolved in a solution of HFIP/MeOH (1:2). Preparation of Liposomes and Supported Bilayers. 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) was purchased from Avanti. Fluorescently labeled lipid N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)-1,2-dihexadecanoyl-sn-glycero-3phosphoethanolamine (NBD-PE) was purchased from Molecular Probes. To prepare unilamellar vesicles (liposomes), lipids and proteins were first mixed in organic solvents (chlorofrom, HFIP, and MeOH). Solvents were dried off under a stream of nitrogen gas, and the mixture was lyophilized and then redissolved in 10 mM phosphate buffer, 100 mM NaCl, pH 7. All vesicles were prepared at 1 mg/mL of POPC, with the desired protein:lipid and donor:acceptor ratios. Protein concentrations are reported here as protein-to-lipid ratios. Samples were vortexed, frozenthawed, and extruded using a 100 nm pore diameter membrane (Avanti). After extrusion, the concentration of the proteins was determined from absorbance measurements, using a Cary UV/ VIS spectrophotometer (Varian). Supported bilayers were prepared by incubating the liposomes with either clean glass or silanized glass surfaces. Experiments on glass surfaces were carried out using an eight-well chambered slide (BD Falcon). Press-to-Seal silicone isolators (Molecular Probes) were used on silanized substrates. Vesicles were incubated in the dark for approximately 1 h. Excess vesicles were removed with an extensive rinse with DI H2O, followed by 10 mM phosphate buffer, 100 mM NaCl, pH 7. Solution FRET. FRET experiments in vesicles were carried out using a Fluorolog fluorometer (Jobin Yvon). The excitation wavelength was set at 450 nm, and emission spectra were collected from 540 to 800 nm (Figure 1A). FRET was measured in liposomes containing known concentrations of donor- and acceptor-labeled proteins. Liposomes containing only donorlabeled proteins (at the same concentration) served as the “no FRET” control. Energy transfer, E, was calculated from measurements of donor intensity at 568 nm, in the absence and presence of the acceptor Cy5:
E (%) ) (ICy3 - ICy3Cy5)/(ICy3) × 100
(1)
where ICy3 and ICy3Cy5 are the intensities of samples containing only Cy3-labeled proteins and samples with both Cy3- and Cy5labeled proteins, respectively. FRAP and Imaging FRET. FRAP and imaging FRET experiments were performed using an Eclipse E600 microscope (Nikon) equipped with a mercury lamp and a SPOT RT camera (Diagnostic Instruments). NBD, Cy3, and Cy5 were excited and observed with the appropriate filter cubes: NBD, excitation 465495 nm, emission 515-555 nm; Cy3, excitation 530-560 nm, emission 573-648 nm; Cy5, excitation 590-650 nm, emission 663-735 nm (Nikon). In the FRAP experiments, an octagonal region (spot) of either 60 or 120 µm diameter was bleached. The spot was set by adjusting the field stop of the microscope using a 10× or a 20× objective. The bleaching time was set to 10 min, and fluorescence recovery was observed with a 10× objective. For the imaging FRET measurements, the liposomes (their solution FRET spectra already collected) were deposited onto the substrate to form the bilayer (see Preparation of Liposomes and Supported Bilayers). The imaging FRET experiments were based on measurements of donor fluorescence before and after acceptor bleaching, and were carried out as follows (see Figure 2): (1) Prior to bleaching, images of Cy3 and Cy5 were captured separately with their respective filter cubes. (2) Using the Cy5 filter, the field stop was lowered and Cy5 was bleached in a small area (∼120 µm in diameter). The decrease in Cy5 intensity was measured during the bleaching process, and the bleaching continued until no significant change in intensity was observed. (3) A Cy3 image was captured after Cy5 bleaching. With Cy5 bleached, no energy transfer from Cy3 to Cy5 occurred, and as a result Cy3 fluorescence increased. The occurrence of FRET was obvious from the appearance of a bright spot in the Cy3
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Figure 2. Method of imaging FRET. (A) “Prebleach” state. When the donor (D) is excited, energy is transferred to the acceptor (A) if the two dyes are in close proximity. (B) The acceptor is bleached. (C). “Postbleach” state. Energy transfer does not occur, and donor fluorescence increases. The FRET % is calculated from the increase in donor fluorescence (eq 2).
Figure 1. (A) Solution FRET measurements in liposomes. The excitation was fixed at 450 nm, such that only Cy3 was directly excited. The emission was scanned from 540 to 800 nm. Solid line: liposomes containing 0.1 mol % Cy3-labeled FGFR3 TM helix (no FRET control). Dashed line: liposomes containing 0.1 mol % Cy3-labeled FGFR3 TM helix and 0.1 mol % Cy5-labeled FGFR3 TM helix. Dotted line: liposomes containing 0.1 mol % Cy5-labeled FGFR3 TM helix. FRET is obvious from the decrease in Cy3 fluorescence (around 570 nm) and the appearance of sensitized Cy5 fluorescence (around 670 nm). The FRET % was calculated from the decrease in Cy3 fluorescence at 568 nm (eq 1). (B) Circular dichroism spectrum of FGFR3 TM helices in liposomes, demonstrating that the proteins are indeed helical. image (Figure 3) in the area where Cy5 had been bleached. Imaging FRET efficiencies were calculated as
E (%) ) (Cy3postbleach - Cy3prebleach)/(Cy3postbleach) × 100, (2) where Cy3postbleach and Cy3prebleach are the measured Cy3 intensities (ImageJ, NIH) after and before the complete photobleaching of Cy5. Background fluorescence in Cy3 intensities was corrected by subtracting the background fluorescence generated from Cy5labeled proteins in the bilayer. Control experiments showed that bleaching of Cy5 did not cause Cy3 bleaching. For these experiments, surface-supported bilayers, containing Cy3-labeled proteins only, were formed. A small area was irradiated using the Cy5 filter for 10-20 min; no Cy3 bleaching was observed.
Results FRAP. Various experimental techniques (neutron reflectivity, SPR, etc.) are used to probe the architecture of
Figure 3. Imaging FRET results. Images of Cy3 and Cy5 are captured before and after Cy5 bleaching. Cy5 is bleached in a small area (∼120 µm) by adjusting the field stop (see the Materials and Methods). The decrease in Cy5 intensity is measured during the bleaching process, and the bleaching continues until no significant change in intensity is observed. The increase in Cy3 fluorescence after Cy5 bleaching is indicative of FRET.
supported bilayers.20-28 As a first step toward the characterization of supported lipid bilayers containing TM helices, we have carried out FRAP studies. FRAP measurements are routinely used to assess lateral mobility in surface-supported lipid bilayers. The FRAP experiment is simple: a small area is bleached, and the fluorescence is then monitored. If the molecules are mobile, the bleached spot gradually disappears due to the lateral diffusion of both bleached and fluorescent molecules. In our experiments, we bleached a small area of the bilayer without a laser, by lowering the field stop for a relatively long time (20) Naumann, R.; Johczyk, A.; Kopp, R.; van Esch, J.; Ringsdorf, H.; Knoll, W.; Graber, P. Angew. Chem., Int. Ed. Engl. 1995, 34, 20562058. (21) Sinner, E. K.; Knoll, W. Curr. Opin. Chem. Biol. 2001, 5, 705711. (22) Schonherr, H.; van Os, M. T.; Forch, R.; Timmons, R. B.; Knoll, W.; Vancso, G. J. Chem. Mater. 2000, 12, 3689-3694. (23) Shen, W. W.; Boxer, S. G.; Knoll, W.; Frank, C. W. Biomacromolecules 2001, 2, 70-79. (24) Duschl, C.; Liley, M.; Lang, H.; Ghandi, A.; Zakeeruddin, S. M.; Stahlberg, H.; Dubochet, J.; Nemetz, A.; Knoll, W.; Vogel, H. Mater. Sci. Eng. 1996, C4, 7-18. (25) Plant, A. L. Langmuir 1993, 9, 2764-2767. (26) Plant, A. L.; Gueguetchkeri, M.; Yap, W. Biophys. J. 1994, 67, 1126-1133. (27) Plant, A. L.; Brigham-Burke, M.; Petrella, E. C.; O’Shannessy, D. J. Anal. Biochem. 1995, 226, 342-348. (28) Plant, A. L. Langmuir 1999, 15, 5128-5135.
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Figure 4. FRAP: Difference in lipid and protein mobility in supported bilayers on glass. Bilayers are composed of POPC doped with NBD-PE (A), and of POPC, NBD-PE, and Cy5-labeled proteins (B and C). FRAP experiments were carried out by bleaching a small area (∼120 µm diameter) for 10 min using the appropriate filter. (A) Recovery of NBD fluorescence in lipid-only bilayers doped with NBD-PE. (B) Partial recovery of NBD fluorescence in mixed protein/lipid bilayers doped with NBD-PE (see also Figure 5). (C) No recovery of Cy5 fluorescence in supported bilayers containing the Cy5-labeled FGFR3 TM helix, showing that the TM helix is immobile (see also Figure 5).
(10 min). Immediately after bleaching, the presence or the lack of mobility can be qualitatively assessed by the visual appearance of the bleached spot. The bleached area has well-defined sharp boundaries if the molecules (lipids or proteins) do not diffuse in the plane of the bilayer during bleaching. The edges of the bleached area appear blurred, however, if the molecules diffuse in the bilayer plane during bleaching. In the latter case, the fluorescence gradually recovers (Figure 4). FRAP measurements of fluorescently labeled proteins in the mixed lipid/protein bilayers on glass revealed that the proteins are immobile. The bleached spot seen in Figure 4C is well defined and does not change its appearance after 30 min (see also Figure 5). This is in contrast to the well-known fact that lipids in supported lipid bilayers on glass are mobile.12,29 Lipid mobility in Figure 4A is evident from the blurred appearance of the bleached area boundaries after bleaching, as well as the complete recovery of fluorescence after 30 min. Since the lack of protein mobility is most probably due to interactions of the proteins with the substrate, we explored the possibility to vary (and control) protein mobility by modifying the substrate. In particular, we used commercially available silanes to modify the surface with various amines, hydroxyls, carboxyls, and thiols. In all studied cases, however, we did not observe protein mobility. We then probed lipid mobility in supported lipid bilayers on the chemically modified glass. We found that some silane modifications decreased and even abolished lipid mobility, while others did not have any effect. For instance, lipid mobility was decreased if the glass slide was silanized with GPDES (exposing a glycidoxy group on the surface), and abolished if the glass slide was silanized with APDES (exposing an amine). From such experiments (not shown), we concluded that, on some substrates, such as ADPES-modified glass, both the (29) van Oudenaarden, A.; Boxer, S. G. Science 1999, 285, 10461048.
Figure 5. NBD recovery after photobleaching in lipid (empty circles) and in mixed lipid/protein (filled circles) bilayers on glass. The ratio of fluorescence intensity in the middle of the bleached spot to peripheral intensity is plotted as a function of recovery time. Bilayers were prepared from liposomes composed of POPC/NBD-PE, with or without 0.5 mol % unlabeled proteins. In 30 min, NBD fluorescence has fully recovered only in the absence of the protein (see the text). Therefore, the presence of proteins in the supported bilayer decreases, but does not abolish, lipid mobility. In contrast, proteins are completely immobile: Cy5 fluorescence (triangles) in bilayers containing 0.1 mol % Cy5-labeled proteins does not recover.
protein and lipid components are immobile while, on other substrates, such as clean glass, the protein is immobile while the lipid is mobile. The latter case is depicted in Figure 4B,C, showing fluorescence recovery after photobleaching of a lipid/protein bilayer supported on glass containing NBD-labeled lipids and Cy5-labeled proteins. It is known that proteins can interact strongly with lipids, and we hypothesized that lipid mobility in the mixed bilayer on glass is reduced, as compared to lipid mobility in lipid-only bilayers. We therefore measured lipid mobility on glass in pure lipid and in mixed protein/lipid bilayers. Figure 5 shows NBD-PE fluorescence recovery after photobleaching in supported lipid bilayers with and without unlabeled proteins. Bilayers were prepared from vesicles composed of POPC doped with NBD-PE, with and
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without unlabeled proteins (0.5 mol %). We see that recovery in the mixed bilayer is not complete after 30 min, in contrast to the lipid-only bilayer. Therefore, lipid mobility is decreased, but not abolished, in the presence of proteins in the bilayer. This decrease in mobility is most probably due to interactions between the lipids and the immobile proteins. The observed difference in mobility of the two components in the mixed supported bilayers prompted us to ask if proteins and lipids tend to phase separate (cluster) in the supported bilayer as a result of different interactions with the substrate. If this is the case, measurements in the supported bilayer systems will not correctly report protein-protein interactions in bilayers. Such phase separation (or substrate-induced protein aggregation) obviously does not occur at the micrometer scale, as evident from the homogeneous fluorescence in Figure 4B,C. Below we present data suggesting that phase separation does not occur at the molecular level, either. In particular, we report that the FRET signal, which is a sensitive reporter of protein aggregation, is identical in liposomes and in surface-supported bilayers. The FRAP and FRET studies, taken together, suggest that the substrate immobilizes the proteins, but does not induce protein dissolution from the lipid matrix. Therefore, protein-protein interactions in the supported bilayer appear to be a frozen “snapshot” of protein-protein interactions in free suspended vesicles. Imaging FRET. FRET is widely used as a spectroscopic “ruler” for detecting molecular interactions in solution, as well as in membranes. It involves the nonradiative transfer of energy from the excited state of a donor molecule to an appropriate acceptor.16,17,30-32 If the two proteins dimerize, the donor and the acceptor will be brought in close contact such that FRET will occur. As a result, the donor fluorescence will decrease and acceptor fluorescence will increase.16,31,32 Different imaging techniques have been developed to measure FRET.33-39 A particular variation of imaging FRET,18,19 based on acceptor photobleaching, has been used by A. K. Kenworthy and M. Edidin16,17 to investigate the organization of GPI-anchored proteins. In this method, the donor fluorescence increases after the acceptor is photobleached, because no energy transfer occurs between donors and bleached acceptors. A detailed explanation of this method is given in the Materials and Methods (see also Figure 2). To verify the imaging FRET methodology, we compared imaging FRET results with those of FRET measured in free suspended liposomes, derived from traditional emission scans (Figure 1A). The FRET % in liposomes was calculated as described in the Materials and Methods. Error bars for the solution experiments were calculated on the basis of errors in absorbance measurements needed to calculate labeled protein concentrations (determined (30) Wu, P.; Brand, L. Anal. Biochem. 1994, 218, 1-13. (31) Clegg, R. M. Curr. Opin. Biotechnol. 1995, 6, 103-110. (32) Clegg, R. M. Fluorescence resonance energy transfer (FRET). In Fluorescence Imaging Spectroscopy and Microscopy; Wang, X. F., Herman, B., Eds.; John Wiley: New York, 1996; pp 179-252. (33) Kubitscheck, U.; Kircheis, M.; Schweitzer-Stenner, R.; Dreybodt, W.; Jovin, T. M.; Pecht, I. Biophys. J. 1991, 60, 307-318. (34) Ju¨rgens, L.; Arndt-Jovin, D.; Pecht, I.; Jovin, T. M. Eur. J. Immunol. 1996, 26, 84-91. (35) Uster, P. S.; Pagano, R. E. J. Cell Biol. 1986, 103, 1221-1234. (36) Kam, Z.; Volberg, T.; Geiger, B. J. Cell Sci. 2000, 108, 10511062. (37) Kindzelskii, A. L.; Xue, W.; Todd, R. F., III.; Petty, H. R. J. Struct. Biol. 1994, 113, 191-198. (38) Kindzelskii, A. L.; Laska, Z. O.; Todd, R. F., III; Petty, H. R. J. Immunol. 1994, 156, 297-309. (39) Oida, T.; Sako, Y.; Kusumi, A. Biophys. J. 1993, 64, 676-685.
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as ∼(10%, on the basis of multiple measurements). After FRET was measured in solution, the liposomes were incubated with the substrate to form the bilayer. Imaging FRET was performed on three different surfaces for each sample: clean glass, glass silanized with GPDES, and glass silanized with APDES. These three substrates were chosen because the lipid mobility on them was different, as discussed above. Images obtained from a typical imaging FRET experiment are presented in Figure 3. Several imaging FRET experiments were performed for each bilayer sample, and the average and standard deviation were calculated. Figure 6 compares solution FRET and imaging FRET results for the three chosen substrates, for four different liposomal samples (A-D). These results show that FRET efficiencies obtained from imaging FRET experiments and solution FRET experiments for each sample are the same within experimental error. Therefore, the substrate does not induce phase separation between lipids and proteins. These results suggest that imaging FRET in supported bilayers could be used as an alternative to solution FRET in liposomes. Furthermore, imaging FRET results for the three substrates, glass, GPDES, and ADPES, are identical within experimental error (and all are identical to the solution FRET results) for each liposomal sample (A-D). Therefore, protein-protein interactions in supported bilayers do not depend on the nature of the substrate. In two control experiments, we measured the imaging FRET signal in bilayers derived from liposomal samples that did not show any solution FRET. In one experiment, imaging FRET experiments were carried out at low protein-to-lipid ratio (0.01 mol %), so that no solution FRET was detected (not shown). In a second experiment, we mixed vesicles containing 0.1 mol % Cy3-labeled proteins with vesicles containing 0.1 mol % Cy5-labeled proteins. These liposomes do not fuse, and therefore do not give solution FRET. In these two cases the Cy3 intensity did not change after Cy5 bleaching (data not shown). Therefore, FRET was negligible in both liposomal solutions and planar supported bilayers. These experiments support the idea that the substrate does not induce protein aggregation. Next, we examined if the imaging FRET results are reproducible. We prepared three different liposomal solutions at identical protein-to-lipid ratios. The imaging FRET was measured, and the results, collected over a 2 month period, are shown in Figure 7. All the experiments were carried out with liposomes containing 0.2 mol % protein, deposited on glass and APDES-coated glass slides. The results are reproducible, suggesting that imaging FRET is a reliable alternative to solution FRET. Data derived from the presented imaging FRET measurements of FGFR3 TM helix interactions in supported bilayers are consistent with the behavior of dimerizing TM helices. If we increase the protein concentration, we expect to observe more dimers and a higher FRET signal.40,41 Figure 8 shows imaging FRET efficiencies measured at three different protein concentrations on clean glass substrates. As expected, the FRET signal increases with the increase in protein-to-lipid ratio. It has been shown that, if the helices form dimers but no higher order aggregates (such that only monomers and dimers are present), FRET depends linearly on the (40) Fisher, L. E.; Engelman, D. M.; Sturgis, J. N. J. Mol. Biol. 1999, 293, 639-651. (41) Fleming, K. G.; Engelman, D. M. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 14340-14344.
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Figure 6. FRET efficiencies measured in liposomal solutions and in planar lipid bilayers on glass, GPDES, and APDES. Liposomes and supported bilayers were formed as described in the Materials and Methods. The solution FRET efficiency was measured in liposomes as shown in Figure 1, prior to their deposition onto the three different surfaces. The imaging FRET efficiency was calculated according to eq 2. Data shown are for four different liposomal samples containing 0.2 mol % FGFR3 TM helix, at 1:2, 2:1, 1:1, and 1:1 Cy3:Cy5 ratios (A, B, C, and D, respectively). The measured FRET efficiencies for each liposomal sample are identical (within experimental error) in solution and on the three surfaces.
Figure 7. Imaging FRET efficiencies from three separate liposomal preparations. The experiments were performed on separate days using freshly prepared samples from different stock solutions of labeled proteins. The lipid concentration was 1 mg/mL, with 0.2 mol % protein. Bilayers were deposited on glass and APDES-modified glass surfaces.
Figure 9. FRET efficiencies measured for 0.2 mol % total peptide concentration as a function of acceptor ratio for bilayers on glass (squares), GPDES (triangles), and APDES (circles). The linear dependence on acceptor mole ratio is characteristic for proteins in a monomer-dimer equilibrium.
Figure 8. Imaging FRET efficiencies as a function of protein concentration. Th FRET efficiency was measured for 0.1, 0.2, and 0.3 mol % FGFR3 TM helix (at a 1:1 Cy3:Cy5 mole ratio) on glass. Errors were determined as described in the Materials and Methods. Results for 0.2 and 0.3 mol % protein are the average from three and two separate liposomal preparations, respectively.
acceptor ratio.42,43 Figure 9 shows FRET efficiencies measured for 0.2 mol % total peptide concentration as a function of acceptor (Cy5) mole ratio in supported bilayers on glass, GPDES, and APDES. For all three surfaces, we
observe straight lines, consistent with monomer-dimer equilibrium. These results provide additional proof that the proteins in the surface-supported bilayers do not form large aggregates as a result of a phase separation between lipids and proteins. Discussion Toward “Membrane Protein Chips”. The incorporation of integral membrane proteins into de novo designed (42) Adair, B. D.; Engelman, D. M. Biochemistry 1994, 33, 55395544. (43) Li, M.; Reddy, L. G.; Bennett, R.; Silva, N. D.; Jones, L. R.; Thomas, D. D. Biophys. J. 1999, 76, 2587-2599.
Transmembrane Helix Interactions in Lipid Bilayers
biomaterials and devices still lies still in the future. The first step toward this goal is the development of membrane protein arrays, a difficult task because the proteins function only within the lipid matrix. The first surfaceimmobilized bilayers with integral membrane proteins were purple membrane patches. Identical orientation of the purple membranes was achieved by an electric field,44 by bispecific antibodies,45 or by chemical modification and selective enzymatic digestion.45 Successful immobilization was monitored by measuring the current generated in response to light. There have also been a few attempts to create surface-immobilized bilayers containing monomeric bacteriorhodopsin by fusion of liposomes containing bacteriorhodopsin to monolayers,46,47 to glass surfaces,47 or to spherical particles.48 In all cases, a photocurrent was observed. Other proteins that have been successfully incorporated into surface-immobilized bilayers are H+ATPase,20,49 the acetylcholine receptor,50 cytochrome b551 and rhodopsin.52 Recently, Fang et al.53,54 demonstrated that G-protein-coupled arrays can be engineered. Here we show that the imaging FRET signal in supported bilayers is comparable to the solution FRET signal in liposomal suspensions. Therefore, the interactions between transmembrane helices in surface-supported bilayers, as inferred from FRET measurements, are the same as in freely suspended liposomal solutions. This is a proof of principle that measurements of TM helix dimerization can be carried out in supported bilayers. An advantage of the imaging FRET measurements in supported bilayers is that the method can be further developed as a high-throughput method. Currently, we perform about eight different experiments on a single microscope slide: we use either eight-well chambered slides (BD Falcon) or Press-to-Seal silicone isolators (Molecular Probes) to create distinct supported bilayer patches. This number could be further increased by patterning the silicon surface or microprinting the supported bilayers. Therefore, the work presented here could be viewed as a first step toward the design of membrane protein chips for parallel measurements of membrane protein interactions. TM Helix Dimerization Studies in Lipid Bilayer Environments. Free energy measurements of TM helix dimerization are needed for understanding the physical principles underlying vital cellular processes.40,41,55-58 For (44) Nicolini, C.; Erokhin, V.; Paddeu, S.; Paternolli, C.; Ram, M. K. Biosens. Bioelectron. 1999, 14, 431-437. (45) Harada, Y.; Yasuda, K.; Nomura, S.; Kajimura, N.; Sasaki, Y. C. Langmuir 1998, 14, 1829-1835. (46) Steinem, C.; Janshoff, A.; Galla, H.-J.; Sieber, M. Chem. Phys. Lipids 1997, 89, 141-152. (47) Puu, G.; Gustafson, I.; Artursson, E.; Ohlsson, P.-Å. Biosens. Bioelectron. 1995, 10, 463-476. (48) Rothe, U.; Aurich, H. Biotechnol. Appl. Biochem. 1989, 11, 1830. (49) Naumann, R.; Jonczyk, A.; Hampel, C.; Ringsdorf, H.; Knoll, W.; Bunjes, N.; Gra¨ber, P. Bioelectrochem. Bioenerg. 1997, 42, 241247. (50) Schmidt, E. K.; Liebermann, T.; Kreiter, M.; Jonczyk, A.; Naumann, R.; Neumann, E.; Kukol, A.; Maelicke, A.; Knoll, W. Biosens. Bioelectron. 1998, 13, 585-591. (51) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400-1414. (52) Heyse, S.; Ernst, O. P.; Dienes, Z.; Hofmann, K. P.; Vogel, H. Biochemistry 1998, 37, 507-522. (53) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124, 2394-2395. (54) Fang, Y.; Frutos, A. G.; Lahiri, J. ChemBioChem 2002, 3, 987991. (55) White, S. H.; Ladokhin, A. S.; Jayasinghe, S.; Hristova, K. J. Biol. Chem. 2001, 276, 32395-32398. (56) White, S. H.; Wimley, W. C. Annu. Rev. Biophys. Biomol. Struct. 1999, 28, 319-365. (57) Popot, J.-L.; Engelman, D. M. Annu. Rev. Biochem. 2000, 69, 881-922.
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instance, the folding of multispanning transmembrane helices into unique three-dimensional structures59 allows membrane proteins to carry out complex biochemical tasks, while lateral dimerization of RTKs is a means of signal transduction across the plasma membrane and, therefore, a key regulator of cell growth and differentiation.4 Further investigations of TM helix dimerization will be aided by the development of adequate assays for membrane protein dimerization in lipid bilayer environments. In our laboratory, we seek to develop novel, reliable, and inexpensive methods for probing TM helix dimerization in lipid bilayers. Here we report a method that uses imaging FRET as a means to quantify TM helix dimerization in surface-supported lipid bilayers. The version of the imaging FRET technique that we usesmeasurements of donor fluorescence prior to and after acceptor photobleachingsis not new:18,19 it has been used previously to assess protein proximity in cells.16,17 Here we develop it further as a quantitative technique to measure FRET due to TM helix dimerization in surfacesupported bilayers. We show that the imaging FRET efficiency, measured in single surface-supported bilayers, is the same as the solution FRET efficiency in liposomes. The presented FRET method has several advantages. Solution FRET measurements of TM helix dimerization are expensive and time-consuming. To measure the FRET efficiency in liposomes containing both donors and acceptors, usually one needs a “no FRET” control. This is a separately prepared lipid sample containing only donorlabeled proteins. Liposome extrusion is required for optical measurements, and in the process some proteins and lipids are lost. Thus, it is crucial that concentrations in both samples are known precisely. Errors in concentration measurements, usually based on sample absorbance, can introduce big errors in FRET calculations. Therefore, it is desirable to develop a method for probing TM helix dimerization in membranes that requires a single sample for FRET measurements. In the presented imaging method, the donor fluorescence after acceptor photobleaching provides the “no FRET” control, such that the FRET signal and the “no FRET” control can be measured simultaneously in one sample. The imaging FRET measurements are less expensive than solution FRET measurements. The method can be adopted and easily set up in any research laboratory that houses a fluorescence microscope (which is less expensive than a fluorometer). Imaging FRET required sample volumes that are about 5 times smaller than typical sample volumes for solution FRET. This also substantially reduces the cost due to the very high cost of the chemically synthesized TM helices. Conclusion We have characterized supported bilayers containing TM helices using FRAP and imaging FRET. We have shown that the FRET signal, a measure of TM helix dimerization, is the same in solution-suspended liposomes and in bilayers on a solid support. Data derived from the imaging FRET measurements of FGFR3 TM helix interactions are consistent with the behavior of dimerizing TM helices. We have demonstrated that imaging FRET in supported bilayers is a reliable and inexpensive method (58) Fleming, K. G.; Ackerman, A. L.; Engelman, D. M. J. Mol. Biol. 1997, 272, 266-275. (59) Lemmon, M. A.; Engelman, D. M. Q. Rev. Biophys. 1994, 27, 157-218.
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for probing the thermodynamics of TM helix interactions in a membrane environment. Acknowledgment. We are grateful to Drs. John Tomich and Takeo Iwamoto for protein synthesis, and to Drs. Anne Kenworthy, Michael Edidin, Wolfgang Knoll, and Eva Sinner for valuable discussions. We acknowledge
Li and Hristova
Genevieve Gallagher’s help with the imaging FRAP experiments. We thank Dr. Peter Searson and his laboratory for access to a microscope in the preliminary stages of the project. This work was supported by NSF Grant MCB 0315663 and Whitaker Grant RG01-0370 to K.H. LA048676L
