Lateral Organization of a Membrane Protein in a ... - ACS Publications

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Langmuir 2006, 22, 5412-5418

Lateral Organization of a Membrane Protein in a Supported Binary Lipid Domain: Direct Observation of the Organization of Bacterial Light-Harvesting Complex 2 by Total Internal Reflection Fluorescence Microscopy Takehisa Dewa,*,†,‡ Ryuta Sugiura,† Yoshiharu Suemori,† Miku Sugimoto,† Toshikazu Takeuchi,† Akito Hiro,† Kouji Iida,§ Alastair T. Gardiner,| Richard J. Cogdell,| and Mamoru Nango*,† Materials Science and Engineering, Nagoya Institute of Technology, Gokiso-cho, Showa-ku, Nagoya 466-8555, Japan, PRESTO, JST, Japan, Nagoya Municipal Industrial Research Institute, 3-4-41 Rokuban-cho, Atsuta-ku, Nagoya 456-0058, Japan, and DiVision of Biochemistry and Molecular Biology, Institute of Biomedical and Life Sciences, UniVersity of Glasgow, UniVersity AVenue, G12 8QQ, U.K. ReceiVed January 28, 2006. In Final Form: March 15, 2006 A unique method is described for directly observing the lateral organization of a membrane protein (bacterial light-harvesting complex LH2) in a supported lipid bilayer using total internal reflection fluorescence (TIRF) microscopy. The supported lipid bilayer consisted of anionic 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1′-glycerol)] (DOPG) and 1,2-distearoly-sn-3-[phospho-rac-(1′-glycerol)] (DSPG) and was formed through the rupture of a giant vesicle on a positively charged coverslip. TIRF microscopy revealed that the bilayer was composed of phase-separated domains. When a suspension of cationic phospholipid (1,2-dioleoyl-sn-glycero-3-ethylphosphocholine: EDOPC) vesicles (∼400 nm in diameter), containing LH2 complexes (EDOPC/LH2 ) 1000/1), was put into contact with the supported lipid bilayer, the cationic vesicles immediately began to fuse and did so specifically with the fluid phase (DOPG-rich domain) of the supported bilayer. Fluorescence from the incorporated LH2 complexes gradually (over ∼20 min) spread from the domain boundary into the gel domain (DSPG-rich domain). Similar diffusion into the domain-structured supported lipid membrane was observed when the fluorescent lipid (1,2-dioleoyl-sn-glycero-3-phosphoethanolamineN-lissamine-rhodamine B sulfonyl: N-Rh-DOPE) was incorporated into the vesicles instead of LH2. These results indicate that vesicles containing LH2 and lipids preferentially fuse with the fluid domain, after which they laterally diffuse into the gel domain. This report describes for first time the lateral organization of a membrane protein, LH2, via vesicle fusion and subsequent lateral diffusion of the LH2 from the fluid to the gel domains in the supported lipid bilayer. The biological implications and applications of the present study are briefly discussed.

Introduction The lateral organization of biological membranes has been widely studied with the goal of understanding the functions of the component lipids and proteins.1 For instance, one version of the heterogeneous lateral distribution of lipids, namely, the socalled lipid domain or “raft”, has been of interest in signal transduction systems in which specific lipids (e.g., sphingolipids and cholesterol) and related membrane proteins form dynamic clusters.1c,2 Heterogeneous segregated arrays of membrane proteins have also been revealed in purple bacterial photosynthetic membranes by atomic force microscopy (AFM).3 Typically, the purple bacterial photosynthetic apparatus consists of reaction center-light harvesting complex 1 (RC-LH1), which is a core complex4 surrounded by several light-harvesting complex 2 * Corresponding authors. (T.D.) E-mail: [email protected]. Tel/ Fax: +81-52-735-5144. (M.N.) E-mail: [email protected]. Tel/Fax: +8152-735-5226. † Nagoya Institute of Technology. ‡ PRESTO, (JST). § Nagoya Municipal Industrial Research Institute. | University of Glasgow. (1) (a) Gennis, R. B. Biomembranes: Molecular Structure and Function; Springer-Verlag: New York, 1989. (b) Yeagle, P. L., Ed. The Structrure of Biological Membranes, 2nd ed.; CRC Press: New York, 2005. (c) Quinn, P. J., Ed. Membrane Dynamics and Domains: Subcellular Biochemistry; Kluwer Academic/Plenum Publishers: New York, 2004. (2) (a) Simons, K.; Ikonen, E. Science 2000, 290, 1721-1726. (b) van Meer, G.; Lisman, Q. J. Biol. Chem. 2002, 277, 25855-25858. (c) Simons, K.; Toomre, D. Nat. ReV. Mol. Cell Biol. 2000, 1, 31-39. (d) Sugahara, M.; Uragami, M.; Regen, S. L. J. Am. Chem. Soc. 2003, 125, 13040-13041.

molecules (LH2).5 Usually, photons are absorbed by LH2, funneled into LH1, and subsequently transferred to the RC, where efficient charge separation takes place. These pigment-protein complexes are ideal model proteins for the present study because their integrity, when incorporated into lipid bilayers, can be monitored spectroscopically and their position can be monitored from their fluorescence, in this case by total internal reflection fluorescence (TIRF) microscopy. This is a powerful tool for observing planar lipid bilayers and biological membranes because it involves the illumination of only a thin region in the evanescent field (∼100 nm) at the coverslip-membrane interface.6,7 In addition, these photosynthetic membrane proteins are potential candidates as components of nanoscale devices.8 Recent (3) (a)Bahatyrova, S.; Frese, R. N.; Siebert, C. A.; Olsen, J. D.; van der Werf, K. O.; van Grondelle, R.; Niederman, R. A.; Bullough, P. A.; Otto, C.; Hunter, C. N. Nature 2004, 430, 1058-1062. (b) Scheuring, S.; Sturgis, J. N.; Prima, V.; Bernadac, A.; Le´vy, D.; Rigaud, J.-L. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 11293-11297. (4) Roszak, A. W.; Howard, T. D.; Southall, J.; Gardiner, A. T.; Law, C. J.; Isaacs, N. W.; Cogdell, R. J. Science 2003, 302, 1969-1972. (5) (a) McDermott, G.; Prince, S. M.; Freer, A. A.; Hawthornthwaite-Lawless, A. M.; Papiz, M. Z.; Cogdell, R. J.; Isaacs, N. W. Nature 1995, 374, 517-521. (b) Papiz, M. Z.; Prince, S. M.; Howard, T.; Cogdell, R. J.; Issacs, N. W. J. Mol. Biol. 2003, 326, 1523-1538. (6) Shaw, J. E.; Slade, A.; Yip, C. M. J. Am. Chem. Soc. 2003, 125, 1183811839. (7) (a) Iino, R.; Koyama, I.; Kusumi, A. Biophys. J. 2001, 80, 2667-2677. (b) Murase, K.; Fujiwara, T.; Umemura, Y.; Suzuki, K.; Iino, R.; Yamashita, H.; Saito, M.; Murakoshi, H.; Ritchie, K.; Kusumi, A. Biophys. J. 2004, 86, 40754093. (8) Markvart, T. Prog. Quantum Electron. 2000, 24, 107-186.

10.1021/la060275d CCC: $33.50 © 2006 American Chemical Society Published on Web 05/10/2006

Direct ObserVation of Membrane Protein Organization

progress in studies on solid-supported membranes9,10 has opened the door to using this technology with these photosynthetic membrane proteins in applications such as molecular devices as well as, more generally, for exploring fundamental functions of membrane proteins. There are two main procedures for making supported membranes that contain membrane proteins. One is to spread reconstituted vesicles on a solid surface, and the other is to use “anchor motifs” in both the membrane components (e.g., modified lipids and engineered proteins) and on the solid surface. Bourdillon’s group has recently demonstrated that inner mitochondrial membranes can be supported on a solid substrate.10c The supramolecular organization of such energy transduction systems, in both respiration and photosynthesis, is intriguing. Biological systems have sophisticated machinery for assembling membrane proteins (e.g., the translocon11). So far, however, it has not been possible to replicate these assemblies artificially. To approach this issue, we have previously described the lateral and transbilayer organization of engineered light-harvesting polypeptides in lipid bilayers.12,13 With respect to the lateral organization, we have demonstrated the lipid-domain selective organization of an engineered LH polypeptide, which suggests that the lipid domain does indeed influence the organization of membrane proteins in such reconstituted systems.12 Such a domain-structured lipid bilayer can be expected to provide a stage for assembling multicomponent membrane proteins in the manner of “the right component in the right place”. Our previous results prompted us to begin to try to achieve this long-term aim by learning how to utilize domain-structured supported lipid bilayers consisting of simple and incompatible binary phospholipid mixtures. As the first step in this process, we propose here a novel approach to study the organization of the LH2 membrane protein in a domain-structured supported lipid bilayer via a vesicle fusion method that is somewhat analogous to processes in biological vesicular transport.11 Previous reports have described the reconstitution of membrane proteins such as bacteriorhodopsin in a giant vesicle (GV)14 or planar membrane15 via vesicle fusion methods; however, to the best of our knowledge, our method is unprecedented. Furthermore, the observed rearrangement of LH2 and lipid molecules in the (9) (a) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105-113. (b) McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A. Biochim. Biophys. Acta 1986, 864, 95-106. (c) Sackmann, E. Science 1996, 271, 43-48. (d) Tanaka, M;, Sackmann, E. Nature 2005, 437, 656-663. (d) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry, 1996, 35, 14773-14781. (e) Groves, J. T.; Ulman, N.; Boxer, S. G. Science 1997, 275, 651-653. (f) Groves, J. T.; Dustin, M. L. J. Immunol. Methods 2003, 278, 19-32. (g) Boxer, S. G. Curr. Opin. Chem. Biol. 2000, 4, 704-709. (h) Tero, R.; Takizawa. M.; Li, Y. -J.; Yamazaki, M.; Urisu, T. Langmuir 2004, 20, 7526-7531. (i) Feigenson, G. W.; Buboltz, J. T. Biophys. J. 2001, 80, 2775-2788. (10) (a) Giess, F.; Friedrich, M. G.; Heberle, J.; Naumann, R. L.; Knoll, W. Biophys. J. 2004, 87, 3213-3220. (b) Ataka, K.; Giess, F.; Knoll, W.; Naumann, R.; H.-Pohlmer, S.; Ritcher, B.; Heberle, J. J. Am. Chem. Soc. 2004, 126, 1619916206. (c) Elie-Caille, C.; Fliniaux, O.; Pantigny, J.; Maziere, J.-C.; Bourdillon, C. Langmuir 2005, 21, 4661-4668. (d) Tero, R.; Misawa, N.; Watanabe, H.; Yamamura, S.; Nambu, S.; Nonogaki, Y; Urisu, T. e-J. Surf. Sci. Nanotech. 2005, 3, 237-243. (e) Ganchev, D. N.; Rijkers, D. T. S.; Snel, M. M. E.; Killian, J. A.; de Kruijff, B. Biochemistry 2004, 43, 14987-14993. (f) Epand, R. M.; Maekawa, S.; Yip, C. M.; Epand, R. F. Biochemistry 2001, 40, 10514-10521. (g) Last, J. A.; Waggoner, T. A.; Sasaki, D. Y. Biophys. J. 2001, 81, 2737-2742. (11) (a) Alberts, B.; Johnson, A.; Lewis, J.; Raff, M.; Roberts, K.; Walter, P. Molecular Biology of the Cell, 4th ed.; Garland Science: New York, 2001; Chapter 12. (b) Hessa, T.; Kim, H.; Bilhlmaier, K.; Lundin, C.; Boekel, J.; Andersson, H.; Nilsson, I.; White, S. H.; von Heijne, G. Nature 2005, 433, 377-381. (12) Dewa, T.; Yoshida, K.; Sugimoto, M.; Sugiura, R.; Nango, M. e-J. Surf. Sci. Nanotech. 2005, 3, 145-150. (13) Dewa, T.; Yamada, T.; Ogawa, M.; Sugimoto, M.; Mizuno, T.; Yoshida, K.; Nakao, Y.; Kondo, M.; Iida, K.; Yamashita, K. Tanaka, T.; Nango, M. Biochemistry 2005, 44, 5129-5139. (14) Kahya, N.; Pe´cheur, E.-I.; de Boeij, W. P.; Wiersma, D. A.; Hoekstra, D. Biophys. J. 2001, 81, 1464-1474. (15) Chanturia, A.; Chernomoridik, L. V.; Zimmerberg, J. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 14423-1428.

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supported lipid bilayer has also not been previously described. A schematic illustration of the process is depicted in Figure 1. The anionic lipid bilayer (either 1,2-dioleoyl-sn-glycero-3[phospho-rac-(1′-glycerol)] (DOPG) in part a or a binary mixture of DOPG and 1,2-distearoly-sn-3-[phospho-rac-(1′-glycerol)] (DSPG) in part b) was supported on a cationically modified coverslip via spontaneous rupture of the GV (A). We chose GV for the formation of the supported lipid bilayer instead of the commonly used small unilamellar vesicle because the domain structure is readily observed by TIRF microscopy. In part b, the domain structure may function as a compartment for depositing multicomponent membrane proteins. The vesicle fusion method is depicted in B, where the cationic proteoliposome plays the role of transporting the LH2 membrane protein to the negatively charged supported lipid bilayers (c and d). TIRF microscopy, which has high sensitivity at the surface of the glass due to the narrow evanescent field (∼100 nm), allows the monitoring of vesicle fusion and subsequent lateral diffusion of the components (lipid and LH2) (c) and the rearrangement of LH2 between the lipid domains (d). In this report, we describe (i) a novel methodology for the incorporation of the LH2 membrane protein into the supported lipid bilayer, (ii) the dynamic process of incorporation and subsequent lateral diffusion of LH2 in the domain-structured supported membrane, (iii) domain-selective organization of LH2, and (iv) lateral rearrangement of the incorporated LH2 into the domain-structured lipid membrane. Experimental Section Unless stated otherwise, all chemicals and reagents were obtained commercially and used without further purification. Phospholipids were purchased from Avanti Polar Lipids, Inc. The LH2 complexes were isolated from membranes of Rhodopseudomonas acidophila strain 10050 and were purified as follows.16 Cells were grown anaerobically in the light, harvested by centrifugation, and suspended in a Tris-HCl solution (20 mM, pH 8.0). Then, upon addition of DNase and MgCl2, the cells were broken by passage through a French press. The photosynthetic membranes were collected from the broken-cell mixture by centrifugation and resuspended in the Tris-HCl solution to an OD at 855 nm of 50. The membranes were then solubilized by the addition of 1% v/v of the detergent N,Ndimethyldodecylamine-N-oxide (LDAO). These solubilized membranes were subjected to sucrose density centrifugation in order to fractionate the sample into LH2 and RC-LH1 complexes. The LH2 fraction was then further purified by ion-exchange chromatography using diethylaminoethyl cellulose (DE52, Whatman). The purified LH2 solution had an absorbance ratio of A858/A280 > 2.5, indicating that it is very pure. A coverslip (Matsunami NEO micro cover glass, 24 × 32 mm, 0.12-0.17 mm in thickness) was chemically cleaned by Piranha treatment (immersion in a mixed solution of H2SO4 (98%)/H2O2 (33%) ) 7/3) for 5 min, dried under a stream of N2, and reacted with 1% (3-aminopropyl)triethoxysilane (APS, Tokyo Kasei Kogyo Co., Ltd) in dry benzene for 4 h at 80 °C. The APS-modified coverslip17 was thoroughly rinsed with methanol and chloroform and then kept in a desiccator in vacuo until use. A GV solution made from either DOPG or DOPG/DSPG and including 1 mol % 1,2dioleoylphosphatidylethanolamine-N-(lissamine-rhodamine B sulfonyl) (N-Rh-DOPE) was prepared in milli-Q water by the electroformation method;18 the thin film of lipids (1 mg) on ITO electrodes was hydrated with 4 mL of milli-Q water under a low(16) PapizA. M. Z.; Hawthornthwaite, M.; Cogdell, R. J.; Woolley, K. J.; Wightman, P. A.; Ferguson, L. A.; Lindsay, J. G. J. Mol. Biol. 1989, 209, 833835. (17) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124, 23942395. (18) (a) Angelova, M. I.; Sole´au, S.; Me´le´ard, P.; Faucon, J. F.; Bothorel, P. Prog. Colloid Polym. Sci. 1992, 89, 127. (b) Angelova, M. I.; Dimitrov, D. S. Faraday Discuss. Chem. Soc. 1986, 81, 303-311.

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Figure 1. Schematic illustration of formation of supported lipid bilayers through the rupture of giant vesicles (A) and subsequent incorporation of LH2 into the supported lipid bilayer as observed by total internal reflection (TIRF) microscopy (B). As illustrated, the process of vesicle fusion and subsequent lateral organization of LH2 is observable by TIRF microscopy. frequency ac field (2 V, 10 Hz) for 2 h at a temperature above the phase-transition temperature of the lipids, ∼25 °C for DOPG (Tm ) ∼ -18 °C) and ∼70 °C for DOPG/DSPG (Tm for DSPG ) 54.5 °C). The GV solution (20 µL) was slowly added to 50 µL of the milli-Q water reservoir deposited on the APS-modified coverslip. After ∼5 min, the supported lipid bilayer formed. Prior to subsequent operations, the remaining GVs on the coverslip were carefully removed by rinsing with milli-Q water several times. A suspension of transporter vesicles consisting of 1,2-dioleoylsn-3-ethylphosphocholine (EDOPC)19 was prepared by extrusion: a dry thin film of 0.2 mg (0.24 µmol) of EDOPC containing 5 mol % N-Rh-DOPE was hydrated with milli-Q water followed by four freeze-thaw cycles and was then extruded through polycarbonate membranes of 0.4, 0.2, and 0.1 µm pore diameter (four extrusions each) at room temperature. A proteoliposome suspension consisting of LH2 and EDOPC was prepared via dialyzing a micellar solution of 0.78 wt % n-octyl β-D-glucopyranoside (OG) in 20 mM TrisHCl buffer (pH 8.0) containing EDOPC and LH2 (EDOPC/LH2 ) (19) (a) MacDonald, R. C.; Ashley, G. W.; Shida, M. M.; Rakhmanova, V. A.; Tarahovsky, Y. S.; Pantazatos, D. P.; Kennedy, M. T.; Pozharski, E. V.; Baker, K. A.; Jones, R. D.; Rosenzweig, H. S.; Choi, K. L.; Qiu, R.; McIntosh, T. J. Biophys. J. 1999, 77, 2612-2629. (b) Rosenzweig, H. S.; Rakhmanova, V. A.; McIntosh, T. J.; MacDonald, R. C. Bioconjugate Chem. 2000, 11, 306-313.

1000/1 (mol/mol)) against a Tris-HCl buffer solution (20 mM, pH 8.0) for 30 h, followed by extrusion through a polycarbonate membrane (0.4 µm pore diameter; four extrusions) at room temperature. In a typical experiment, vesicle fusion was carried out as follows: the proteoliposome solution (0.5 µM of LH2, 20 µL) was carefully placed in a 30 µL milli-Q water reservoir that had been deposited on the supported lipid bilayer (at room temperature). Fluorescence microscopy observation was performed with an objective-type TIRF microscope, TE-2000U (Nikon), equipped with an oil-immersion objective lens (Plan Apo 100×H; numerical aperture ) 1.45, Nikon), a laser with λ ) 532 nm (25 mW, Crystalaser), a cooled CCD camera (ORCA-ER: HAMAMATSU), and AQUACOSMOS imaging analysis software. The filter set used was a 575 nm dichroic mirror/barrier filter (590 nm long pass) for experiments involving the fluorescent lipid N-Rh-DOPE. For observation of the LH2 complex, with a fluorescence band in the near-infrared region, an additional band-pass filter, 765-855 nm, was used. Fluorescence recovery after photobleaching (FRAP) measurement was carried out with a confocal laser scanning microscope (either FV1000 (Olympus) or C1 (Nikon)). A bleachable fluorescent probe, 1,2dioleoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-ly) (N-NBD-DOPE), was incorporated at 1 mol % in the supported lipid bilayer.

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Figure 2. Total internal reflection fluorescence (TIRF) microscopy images of various supported lipid bilayers and fluorescence intensity profiles along the dashed lines in the images. The supported lipid bilayers contained 1 mol % of the fluorescent lipid probe, N-Rh-DOPE. (a) DOPG; (b, c) DOPG/DSPG (1/1 mol/mol). The bright area represents the fluid DOPG phase. The dark areas in b and c correspond to DSPG-rich domains. The domain size depends on the cooling rate applied during the preparation of the GV. The first cooling after electroformation generated small domains (∼1-5 µm in the size) (b). Slow cooling generated larger domains (c). Images are 30 × 30 µm. Fluorescence intensity profiles of the dashed lines in these images are shown in d-f. Arrows in images (b, c) and fluorescence profiles (e-f) indicate domain boundaries.

Results and Discussion Formation of a Supported Lipid Bilayer on the APSModified Coverslip. A planar lipid bilayer supported by the APS-modified coverslip formed spontaneously when a suspension of GV composed of DOPG or DOPG/DSPG was loaded onto the coverslip.20 In contrast, a GV suspension consisting of the zwitterionic lipid 1,2-dioleoyl-sn-3-phosphocholine does not form such a supported membrane on the APS-modified coverslip, suggesting that electrostatic interaction between the negatively charged bilayer and the cationic surface of the coverslip is responsible for the formation of the supported bilayer. When the DOPG- or DOPG/DSPG-supported lipid bilayer incorporating N-Rh-DOPE was immersed in a buffer solution (20 mM TrisHCl, 100 mM NaCl, pH 7.5), the dissociation of the lipid bilayer from the coverslip was detected by the absorption spectrum of N-Rh-DOPE in the solution. This suggests an electrostatic interaction between the bilayer and support. TIRF microscopy revealed the process whereby the supported lipid bilayer formed: upon deposition of the vesicle suspension onto the coverslip, the GVs contacting the cationic surface were observed to rupture immediately and form a planar bilayer. Multiple vesicle ruptures gave a surface fully covered by a lipid bilayer within ∼5 min. GVs that approached a patch of supported bilayer that had already formed were repelled from the surface of the patch without adhesion or multilayer formation (data not shown). These observations on the process of formation of the supported membrane suggest that it consists predominantly of a single bilayer. Figure 2a-c shows the TIRF microscopy images of the DOPG(a) and DOPG/DSPG-supported membranes (b, c). The bright (fluorescent) and dark areas correspond to fluid (rich in DOPG) and gel (rich in DSPG) domains, respectively.21 The image of (20) The surface smoothness of the cleaned and APS-modified coverslips was evaluated by AFM. In the range of 3 µm of line profile, asperities were less than 0.7 for the cleaned bare coverslip and 10 nm for APS-modified coverslip, respectively. (21) The fluorescent lipid, N-Rh-DOPE, is preferentially incorporated into the fluidic DOPG domain.

the DOPG-supported bilayer (a) suggests a homogeneous structure. The fluorescence intensity profile of a line through the image (d) is smooth, indicating the absence of multilamellar regions. The slightly curved profile is characteristic of evanescent illumination.22 In contrast to the homogeneous texture of the DOPG-supported bilayer (a), phase-separated domain structures were clearly observed for the phase-separating binary lipid system, DOPG/DSPG (b, c). The size of the domains depended on the cooling rate during GV preparation: fast cooling from 70 to 25 °C at 3 °C/min produced small DSPG-rich domains (b) and slow cooling at 0.3 °C/min generated large domains (c). The intensity profile along a line through the domain structure is shown in e and f. The domain in the latter image exhibits a clearer domain boundary than that of the former image, indicating that slowly grown domains are more extensively phase-separated. Fluidity of the Supported Lipid Bilayer. The fluid character of a supported membrane is essential for a model of biological membranes. The DOPG-supported membrane containing the fluorescent probe N-NBD-DOPE was subjected to fluorescence recovery after photobleaching (FRAP) experiments (Figure 3). The fluorescence of the bleached area, whose radius was 1.4 µm, recovered by more than 90% in 10 min, indicating that the supported lipid possesses lateral mobility. The diffusion coefficient, D, is related to the half-life of the recovery according to D ) r2/4τ,9d where r is the radius of the bleaching spot and τ is the time for half-maximal recovery. The experimentally determined diffusion coefficient was in the range of 0.3-0.5 µm2/s and is about an order of magnitude smaller than that reported for other supported, tethered membranes.10c,23 The smaller diffusivity in the present experiments is presumed to be predominantly due to the electrostatic interaction between the lipids and the APS-modified surface. Although the supported lipid bilayer may well be influenced by the electrostatic (22) Because the intensity profile of the laser cross section has a Gaussian distribution, the intensity of the evanescent illumination can be approximately regarded as this distribution. (23) Johnson, J. M.; Ha, T.; Chu, S.; Boxer, S. G. Biophys. J. 2002, 83, 33713379.

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Figure 3. Fluorescence photobleaching of DOPG-supported lipid bilayers containing 1 mol % N-NBD-DOPE. The frame size is 30 × 30 µm, and the diameter of the photobleaching area is 2.8 µm (a) before photobleaching and (b) just after bleaching. Images in c-f represent the fluorescence recovery after 0.7 (c), 1.4 (d), 3.0 (e), and 10.0 s (f).

Figure 4. Sequential snapshots of the incorporation of LH2 via fusion of proteoliposomes with a DOPG-supported bilayer. Fluorescence from the incorporated LH2 was detected by TIRF microscopy (a) before fusion, (b) at the moment of contact of the proteoliposome on the supported lipid bilayer, (c) 360 ms after the touch down, and (d) after 1.44 s. Images are 6 × 6 µm.

Dewa et al.

Figure 5. Time courses of fluorescence decay for LH2 (O) and N-Rh-DOPE (0) for vesicle fusion and probe diffusion within the DOPG-supported membrane. The fluorescence intensity, which was monitored with a CCD camera on the TIRF microscope, was normalized according to Irel ) (I - I′)/(I0 - I′), where I and I′ represent the observed and background intensity, and I0 corresponds to t ) 0.

Figure 6. NIR absorption spectra of LH2 that had been incorporated into a DOPG-supported membrane (-) and dissolved in a buffer solution (20 mM Tris-HCl, pH 8.0, containing 0.1 wt % LDAO) (- -). The spectrum was acquired from a stack of five coverslips of an LH2-incorporated supported membrane.

interactions, the magnitude of the diffusion coefficient itself is close to that of a cell membrane,7b,24 which possesses transmembrane asymmetry due to attachment to the cytoskeleton and other anchor proteins. We believe, therefore, that the supported lipid bilayer described here represents a valid model for biomembranes. Cationic Vesicle Fusion with the Anionic Supported Lipid Bilayer. The incorporation of LH2 into the negatively charged supported lipid bilayer via vesicle fusion of positively charged proteoliposomes (EDOPC/LH2 ) 1000/1 (mol/mol), ∼0.4 µm in diameter) was successfully captured by TIRF microscopy. Figure 4 shows sequential snapshots of such an event. On the DOPG-supported bilayer (the dark area in a), the fluorescent spot (LH2 in the proteoliposome) touches down (b); subsequently, the spot diffuses within the membrane (c, d) in a few seconds. Another experiment involving N-Rh-DOPE instead of LH2 in an EDOPC vesicle (∼0.4 µm in diameter) underwent a similar process. These experiments reveal the vesicle fusion and subsequent lateral diffusion of LH2 and the lipid (N-Rh-DOPE) within the supported lipid bilayer. When vesicles were prepared from EDOPC/DOPG (1/1) or DOPG, the frequency of fusion was lower than that for EDOPC vesicles, suggesting that vesicle fusion depends on electrostatic interaction. In Figure 5 are shown time courses of fluorescence intensity for vesicle fusion and subsequent lateral diffusion. When the fluorescent vesicle (including N-Rh-DOPE or LH2) touches down on the surface of the supported lipid bilayer, a jump in the fluorescence intensity was detected by the CCD camera of the TIRF microscope. The fluorescence decayed in ∼7 and ∼30 s for vesicles containing N-Rh-DOPE and LH2, respectively, and was accompanied by the diffusion of the fluorescent molecules

within the supported bilayer as shown in Figure 4. The time and fluorescence intensity were normalized as follows: the abrupt rise in intensity representing the contact of the vesicle or proteoliposome with the surface (i.e., the situation in Figure 4b) was defined as t ) 0, and the relative fluorescence intensity was defined as I0 ) 1. As mentioned above, the decay of fluorescence for the fusion of vesicles containing N-Rh-DOPE was faster (∼7 s) than that for the LH2-proteoliposome system (∼30 s). The difference in the diffusivity is reasonable in terms of the size of the two fluorescent molecules, the lipid being much smaller than the protein (LH2 is a transmembrane hollow cylinder of ca. 7 nm diameter and ca. 5 nm height).5 The height of LH2 is almost same as the bilayer thickness, so it is reasonable that LH2 diffuses within the supported lipid bilayer without strong steric friction with the substrate. Figure 6 shows the near-IR absorption spectrum of LH2 incorporated into the DOPG-supported membrane (solid line). The spectrum is almost identical to that of the original LH2 sample dissolved in the Tris-HCl (20 mM, pH 8.2) solution containing 0.1% v/v LDAO (dashed line); B850 and B800 appeared at 858 and 802 nm, respectively. The absorbance ratio of B850/B800 ) 1.3 for the incorporated LH2 is in close agreement with the literature value, 1.49.25 This indicates that the method described above is able to carry the LH2 complexes into the supported lipid bilayer without significant denaturation. Domain-Selective Incorporation and Rearrangement of LH2 within the Domain Structure of the Supported Lipid Bilayer. When the vesicle transport procedure, as described above, was applied to a domain-structured supported membrane, domainselective incorporation of LH2 was observed. Figure 7 shows the process. Figure 7a is the image of a phase-separated domain (the same as shown in Figure 2c). The bright and dark areas

(24) Sonnleitner, A.; Schu¨tz, G. J.; Schmidt, Th. Biophys. J. 1999, 77, 26382642.

(25) Georgakopoulou, S.; Frese, R. N.; Johnson, E.; Koolhaas, C.; Cogdell, R. J.; van Grondelle, R.; van der Zwan, G. Biophys. J. 2002, 82, 2184-2197.

Direct ObserVation of Membrane Protein Organization

Figure 7. Incorporation of LH2 into the fluid domain and its subsequent reorganization via lateral diffusion into the gel domain as detected by TIRF microscopy. (a) DOPG-fluid domain (bright area due to incorporated N-Rh-DOPE) and DSPG-gel domain (dark area); (b) image after photobleaching of (a); (c)-(f) snapshot of LH2 incorporation via vesicle fusion and its subsequent lateral diffusion into the gel domain. The images in c-f were taken after the addition of LH2/EDOPC proteoliposomes after 5 (c), 10 (d), 15 (e), and 20 min (f). The bright area is due to fluorescence from LH2. The scale bar represents 10 µm. The change in the fluorescence intensity profiles along the dashed line crossing the domain boundary indicated in c during the time range of 5-20 min is shown in Figure S1 of Supporting Information.

correspond to DOPG-rich (containing N-Rh-DOPE) and DSPGrich domains, respectively. N-Rh-DOPE incorporated within the DOPG-rich fluid domain was photobleached (b). Then, an aliquot of an LH2-proteoliposome suspension was loaded onto the supported membrane. Vesicle fusion was observed to occur selectively within the fluid domain. As fusion progressed (c), the same gel domain structure as that shown in part a appeared. This observation indicates that LH2 is selectively incorporated into the fluid domain. Over a time period of ∼10 min, the gel domain did not change shape (d); however, LH2 subsequently began to cross the domain boundary (e: after 15 min) and gradually invaded the interior of the domain (f: after 20 min). The change in the fluorescence intensity profiles over the time range of 5-20 min along the dashed line in the image of Figure 7c is shown in Figure S1 in Supporting Information. During these stages, vesicle fusion continued until the proteoliposomes were consumed. Such a rearrangement of LH2 in the gel domain suggests that LH2 became incorporated into the fluid phase and then laterally diffused, even into the gel domain. The possibility of the vesicle fusion process on the gel domain can be excluded because although we observed only adhesion to the gel domain we never observed merging of the proteoliposome probe into the gel domain (the small, bright spots seen in e and f). Eventually, the gel domain was fully invaded by LH2 molecules, but during this process, the domain shape did not change. When a lower concentration of the EDOPC/LH2 proteoliposome suspension (∼0.01 µM LH2) was applied, such an intrusion was not observed. Thus, the rearrangement of LH2 into the gel domain seems to depend on the concentration of LH2 in the fluid domain. In other

Langmuir, Vol. 22, No. 12, 2006 5417

words, the distribution of LH2 between the fluid and gel domains depends on conditions that are controllable. A similar intrusion of LH2 complexes into the gel domain was observed when we used a surfactant instead of cationic liposomes. The addition of LH2 (0.5 µM) dissolved in a solution of 0.1% OG onto the domain-containing bilayer resulted in the initial incorporation of LH2 into a fluid domain, followed by intrusion into the adjacent gel domain across the domain boundary. In sharp contrast to the vesicle fusion system, this intrusion happened very fast and was completed within 1 min. That such a phenomenon might occur is not unreasonable because the surfactant makes the lipid bilayer fragile;26 that is, the surfactant may loosen the packing of the bilayer in both fluid and gel domains, which results in rapid incorporation and intrusion processes. As another control experiment, the EDOPC vesicle incorporating fluorescent lipid N-Rh-DOPE instead of LH2 was examined for such lateral diffusion across the phase boundary. Similar fusion and crossing of the boundary were confirmed. (Figure S2 is a movie file in Supporting Information). Biological Implication and Applications. We have demonstrated here that a bilayer vesicle can transport the membrane protein, LH2, specifically into the fluid domain (as the target part) of a supported bilayer via vesicle fusion. We expect that this methodology will also be applicable to the investigation of more complex assembly processes in supported lipid bilayers such as mixtures of LH2 and RC-LH1 complexes. The fluorescence bands of LH2 and RC-LH1 are distinguishable; therefore, TIRF microscopy combined with this assembly method could be used to investigate the lateral organization and diffusion of mixtures of both LH2 and RC-LH1 in the same lipid bilayer. The physicochemical factors that may affect the relative distribution of these complexes can then be studied. Additional applications of the method include assays to study intramembrane protein-protein and lipid-protein interactions (e.g., signaling proteins such as cell surface receptors). In domain-separated lipid bilayers that are composed of liquid ordered (lo)/liquid disordered (ld) phases, the observation of protein/lipid rearrangement is of interest because of the functional importance of their dynamics.27 The cationic phospholipid EDOPC was first used as a nonviral gene carrier,19 but bilayer vesicles of this lipid undergo fusion with anionic GVs.28 The physical properties of EDOPC are similar to those of the naturally occurring phosphatidylcholine,19a and this may be one of the reasons that the vesicle fusion with the supported membrane occurred with good reproducibility. A disadvantage of previous fusion methods, as has been pointed out recently, is the limited amount of membrane protein that can be incorporated.29 Our present method is simple but possesses several key advantages; namely, (i) sequential incorporation of multicomponent membrane proteins is feasible, (ii) use of a domain-structured lipid bilayer makes the functional organization of membrane proteins possible, and (iii) rearrangement and interaction of membrane proteins and lipid can be readily detected. This study, therefore, provides novel experimental methodology for investigating the lateral organization of membrane proteins (26) Nomura, F.; Nagata, M.; Inaba, T.; Hiramatsu, H.; Hotani, H.; Takiguchi, K. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 2340-2345. (27) (a) Kahya, N; Brown, D. A.; Schwille, P. Biochemistry 2005, 44, 74797489. (b) Baumgart, T.; Hess, S. T.; Webb, W. W. Nature 2003, 425, 821-824. (c) Dietrich, C.; Bagatolli, L. A.; Volovyk, N. Z.; Thompson, N. L. Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 1417-1428. (d) Feigerson, G. W.; Buboltz, J. T. Biophys. J. 2001, 80, 2775-2788. (28) (a) Lei, G.; MacDonald, R. C. Biophys. J. 2003, 85, 1585-1599. (b) Pantazatos, D. P.; MacDonald, R. C. J. Membr. Biol. 1999, 170, 27-38. (29) Girard, P.; Pe´cre´aux, J.; Lenoir, G.; Falson, P.; Rigaud, J.-L.; Bassereau, P. Biophys. J. 2004, 87, 419-429.

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and may provide a way to assemble ordered arrays of membrane proteins for use in nanoscale biological devices.

the BBSRC (R.J.C.). M.N thanks professor Robert C. MacDonald for critical comments and discussions.

Acknowledgment. This work is supported by PRESTO (Japan Science and Technology Agency), a Grant-in-Aid for Scientific Research on Priority Areas (417) from the Ministry of Education, Culture, Sports, Science and Technology, NEDO International Joint Grant, Japan, the NITECH 21st Century COE Program “World Ceramics Center for Environmental Harmony”, and also

Supporting Information Available: Change in the fluorescence intensity profiles along the line crossing the domain boundary and a movie of vesicle fusion with the fluid domain and subsequent lateral diffusion into the gel domain. This material is available free of charge via the Internet at http://pubs.acs.org. LA060275D