Formation of Tethered and Streptavidin-Supported Lipid Bilayers on a

Langmuir 2002, 18, 3263-3272

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Formation of Tethered and Streptavidin-Supported Lipid Bilayers on a Microporous Electrode for the Reconstitution of Membranes of Large Surface Area Vanessa Proux-Delrouyre,† Ce´line Elie,† Jean-Marc Laval,† Jacques Moiroux,‡ and Christian Bourdillon*,† Laboratoire de Technologie Enzymatique, UMR du CNRS No. 6022, Universite´ de Technologie de Compie` gne, B.P. 20529, 60205 Compie` gne Cedex, France, and Laboratoire d’Electrochimie Mole´ culaire, UMR du CNRS Nο. 7591, Universite´ Paris 7sDenis Diderot, 75251 Paris Cedex 05, France Received October 23, 2001. In Final Form: January 25, 2002 A new approach for the self-assembly of supported and tethered lipid membranes of large surface area is proposed. The template is a microporous electrode made by anodic etching of aluminum and covered with a monolayer of streptavidin. We show that spontaneous fusion of biotinylated lipid vesicles on the affinity layer is a slow process despite abundant accumulation of lipid material at the template surface. To increase dramatically the efficiency of the self-assembly, fast fusion is provoked with the help of a fusogen solution of poly(ethylene glycol). The extent of fusion is assessed by electrochemical monitoring of the long-range lateral mobility of ubiquinone (coenzyme Q10) in the supported bilayer. Finally, the geometrical characterization of the honeycomb structure at key steps of the self-assembly procedure is performed by electrochemical measurement of the porosity. As expected, the formation of the supported bilayer causes a decrease in the apparent inner diameter of the pores. It is expected that the type of supported lipid membrane built according to the present approach can be adequate for the incorporation of transmembrane proteins in structures that would mimic the membrane stacking found in chloroplasts or mitochondria.

Introduction Electron microscopy photographs of organelles from eukaryotic animal or plant cells reveal regions of closely stacked membranes exhibiting a high concentration of membrane components. For example, there is the equivalent of about 2 × 105 cm2 of inner membranes in 1 cm3 of mitochondria.1 This explains a large part of the high dynamic efficiency of the electron-transfer chains embedded in the cellular structures. The reconstitution of similar structures mimicking the functionality of nanometer-sized biological components, in the intermediate range between 5 and 500 nm, is an interesting challenge either for a better understanding of subcellular behaviors or for future technological applications of biological functions.2 Supported membranes3 represent a promising strategy in this area of research as the use of a solid substrate increases the stability of the bilayer and opens many * To whom correspondence may be addressed. E-mail: [email protected]. Tel.: (33) 3 44 23 44 05. Fax: (33) 3 44 20 39 10. † Laboratoire de Technologie Enzymatique, UMR du CNRS No. 6022, Universite´ de Technologie de Compie`gne. ‡ Laboratoire d’Electrochimie Mole ´ culaire, UMR du CNRS No. 7591, Universite´ Paris 7sDenis Diderot. (1) (a) Hackenbrock, C. R.; Chazotte, B.; Gupte, S. S. J. Bioenerg. Biomemb. 1986, 18, 331-368. (2) (a) Muller, W.; Ringsdorf, H.; Rump, E.; Wildburg, G.; Zhang, X.; Angermaier, L.; Knoll, W.; Liley, M.; Spinke, J. Science 1993, 262, 17061708. (b) Cullison, J. K.; Hawkridge, F. M. Langmuir 1994, 10, 877882. (c) Cornell, B. A.; Braach-Maksvytis, L. G.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wiesczorek, L.; Pace, R. J. Nature 1997, 387, 580583. (d) Heyse, S.; Ernst, O. P.; Dienes, Z.; Hofmann, K. P.; Vogel, H. Biochemistry 1998, 37, 507-522. (3) Recent reviews in: (a) Sackmann, E. Science 1996, 271, 43-48. (b) Wong, J. Y.; Majewski, J.; Seitz, M.; Park, C. K.; Israelachvili, J. N.; Smith, G. S. Biophys. J. 1999, 77, 1445-1457. (c) Plant, A. L. Langmuir 1999, 15, 5128-5135. (d) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400-1414.

possibilities for its characterization. Since the pioneering works on glass surfaces by the McConnell group,4 several successive approaches have demonstrated the interest of a polymer cushion to separate the membrane from the solid support.5,6 Reactions in membranes depend on lateral motion and the fluid dynamic properties of all membrane components, and a soft hydrophilic cushion would alleviate most of the mobility problems encountered when bare or alkylated supports are used.3d In the literature, the planar geometry of the support has been generally considered as essential for the formation and the characterization of supported bilayers. This constraint resulted either from the nature of the bilayer assembly procedures, for example, by Langmuir-Blodgett deposition,3d,5b or more frequently from requirements related to the techniques of bilayer characterization. Numerous methods of control have been developed on flat surfaces indeed: electrical impedance,7 surface plasmon resonance (SPR),2c,5a fluorescence microscopy,3d,4 atomic (4) (a) Brian, A. A.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 6159-6163. (b) Watts, T. H.; Brian, A. A.; Kappler, J. W.; Marrack, P.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 7564-7568. (c) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105-113. (d) Boxer, S. G. Curr. Opin. Chem. Biol. 2000, 4, 704-709. (5) (a) Spinke, J.; Yang, J.; Wolf, H.; Liley, M.; Ringsdorf, H.; Knoll, W. Biophys. J. 1992, 63, 1667-1671. (b) Ku¨hner, M.; Tampe´, R.; Sackmann, E. Biophys. J. 1994, 67, 217-226. (c) Williams, L. M.; Evans, S. D.; Flynn, T. M.; Marsh, A.; Knowles, P. F.; Bushby, R. J.; Boden, N. Langmuir 1997, 13, 751-757. (d) Zhang, L.; Longo, M. L.; Stroeve, P. Langmuir 2000, 16, 5093-5099. (6) (a) Seitz, M.; Wong, J. Y.; Park, C. K.; Alcantar, N. A.; Israelachvili, J. N. Thin Solid Film 1998, 327, 767-771. (b) Seitz, M.; Ter-Ovanesyan, E.; Hausch, M.; Park, C. K.; Zasadzinski, J. A.; Zentel, R.; Israelachvili, J. N. Langmuir 2000, 16, 6067-6070. (7) (a) Steinem, C.; Janshoff, A.; Ulrich, W. P.; Sieber, M.; Galla, H. J. Biochim. Biophys. Acta 1996, 1279, 169-180. (b) Wiegand, G.; Neumaier, K. R.; Sackmann, E. Rev. Sci. Instrum. 2000, 71, 23092320.

10.1021/la011585t CCC: $22.00 © 2002 American Chemical Society Published on Web 03/12/2002

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force microscopy (AFM),8 photobleaching (FRAP),3d,4 and neutron reflectivity.6 Our approach consists of building supported bilayers of large surfaces, with compactness similar to that found, for example, in invaginated inner membranes of mitochondria. We have chosen to proceed with a support made of porous aluminum oxide in such a way that a high surface area/volume ratio can be artificially achieved. The honeycomb disposition of anodic porous alumina is a very convenient template for the development of nanometersized structures.9 It has been already proposed as a support for the directional assembly of amphiphilic layers.10 Our group has developed hybrid lipid layers, lipid monolayers on alkylated aluminum oxide, supported on the same template for the study of lateral diffusion of isoprenic quinones,11 and recent preliminary results have shown that this template can be used successfully for the construction of streptavidin-supported bilayers.12 The method is based on the fabrication of a “microporous electrode” made of a thin porous aluminum oxide film coated with gold by vapor deposition on one side as described in Figure 1. Porous alumina is the template of the supported bilayer, and electrochemical measurements can be carried out at the gold electrode located at the bottom of each pore. Fusion of vesicles is the only possible method of bilayer formation in this geometry. Different variants of fusion of vesicles on a streptavidin sublayer are tested, and a two-step method is finally put forward. Two complementary driving forces are thus involved successively in the assembly of the bilayer as indicated in Figure 2. In the first step, biotinylated small unilamellar vesicles (SUV) are accumulated in the pores due to their interaction with a streptavidin sublayer. In the second step, the fusion of the anchored vesicles and the formation of the bilayer are triggered by treatment with a watersoluble fusogen, poly(ethylene glycol) (PEG), that has been frequently used for the fusion of vesicles in solution.13 The expected final structure is a continuous bilayer covering the whole porous structure, cushioned on the streptavidin-covered surface, and tethered to the support by linkage of the biotinylated lipids with the streptavidin molecules. One critical question in the formation of supported bilayers is the quality of the lateral fusion between vesicles. To assess this lateral fusion, we develop two electrochemical approaches taking advantage of the specific geometry of the microporous electrode. In the first one the lateral diffusion of ubiquinone (coenzyme Q10), a hydrophobic natural electron carrier, is monitored electrochemically. The electrochemically active ubiquinone is strictly water insoluble; therefore the Q10 molecules are incorporated only within the hydrophobic core of the bilayer and can reach the gold electrode only by diffusion within this medium. The fusion of vesicles (8) (a) Singh, S.; Keller, D. J. Biophys. J. 1991, 60, 1401-1410. (b) Jass, J.; Tjarnhage, T.; Puu, G. Biophys. J. 2000, 79, 3153-3163. (9) (a) Masuda, H.; Fukuda, K. Science 1995, 268, 1466-1469. (b) Hennesthal, C.; Steinem, C. J. Am. Chem. Soc. 2000, 122, 8085-8086. (10) Miller, C. J.; Majda, M. J. Electroanal. Chem. 1986, 207, 49-53. (b) Majda, M. Kinetics and catalysis in microheterogeneous systems; Gratzel, M., Kalyanasundaram, K., Eds.; Marcel Dekker: New York, 1991; pp 227-272. (11) (a) Marchal, D.; Boireau, W.; Laval, J.-M.; Moiroux, J.; Bourdillon, C. Biophys. J. 1997, 72, 2679-2688. (b) Marchal, D.; Boireau, W.; Laval, J.-M.; Moiroux, J.; Bourdillon, C. Biophys. J. 1998, 74, 1937-1948. (12) Proux-Delrouyre, V.; Laval, J.-M.; Bourdillon, C. J. Am. Chem. Soc. 2001, 123, 9176-9177. (13) Recent reviews in: Arnold, K. Structure and dynamics of membranes; Lipowsky, R., Sackmann, E., Eds.; Elsevier: Amsterdam, 1996; pp 903-957. (b) Lentz, B. R.; McIntyre, G. F.; Parks, D. J.; Yates, J. C.; Massenburg, D. Biochemistry 1992, 31, 2643-2653.

Proux-Delrouyre et al.

Figure 1. Schematic longitudinal section (A) and front view (B) of the microporous electrode. The honeycomb structure was produced by aluminum anodic corrosion, and the pores were closed by gold vapor deposition under vacuum to produce an electrode at the bottom of each pore. The continuous lipid bilayer covering the inner walls of the pores is built on a cushion of streptavidin molecules. (C) Schematic representation of the molecular assembly in which the supported lipid bilayer is tethered to immobilized streptavidin in the region surrounding the bottom of a pore. Note that the presence of the OM layer attached to the gold surface allows for the continuity of the hydrophobic structure.

Figure 2. PEG-triggered fusion of vesicles inside the microporous electrode. Longitudinal section of a pore before and after PEG treatment. Upper sketch: attachment and accumulation of biotinylated SUV on the streptavidin sublayer. Lower sketch: vesicle rupture and fusion lead to the formation of the supported bilayer on the inner walls of the pores.

containing small amounts of Q10 into a continuous bilayer results in an increase of the available surface for the diffusion of the quinone in two dimensions. Thus the

Tethered and Streptavidin-Supported Lipid Bilayers

electrochemical measurement of the amount of Q10 able to reach the gold electrode by lateral diffusion reflects the formation and the continuity of the bilayer. The second approach is essentially geometrical. The pore diameter being not much larger than the manipulated lipidic structures, the apparent diameter of the pore can be significantly reduced if vesicles are anchored along the pore or if a continuous bilayer covers the inner cylinder. The ensuing alteration of the apparent porosity of the structure can be followed by measurement of the rate of diffusion through the pores of a water-soluble electrochemically active molecule using a rotating disk electrode (RDE) and a modified Koutecky-Levich data analysis.10,14 The comparison with a geometrical model of partially blocked pores gives information on the fusion of vesicles. In the present paper, we optimize the parameters for a routine formation of supported bilayers of large surface area in order to demonstrate that the use of the microporous structure offers new opportunities for the study of supported membrane assembly. Materials and Methods Materials. L-R-Phosphatidylcholinedimyristoyl (DMPC) synthetic, l-R-phosphatidyl-ethanolamine-dimyristoyl (DMPE) synthetic, L-R-phosphatidylcholine (eggPC) type XVI-E from egg yolk, and ubiquinone (Q10) were purchased from Sigma (St Quentin Fallavier, France). Biotin-amidocaproic acid 3-sulfo-N-hydroxysuccinimide ester (NHS-lc-biotin), streptavidin, n-octyl β-Dglucopyranoside (OG), and poly(ethylene glycol) 8000 (PEG), average molecular weight 8000 g mol-1, were also from Sigma. 1,2-Dipalmitoyl-sn-glycero-3-phosphoethanolamin-N-biotinyl (biotinylated DPPE) was from Avanti polar-lipid. Aminopropyldimethylethoxysilane (ADMS), aminoethanethiol, octadecyl mercaptan (OM), and ruthenium(III) hexamine chloride were from (Aldrich, Strasbourg, France). Aluminum foil, 1 mm thick (Al 99.95%) was from Merck (Darmstadt, Germany). Organic solvents were HPLC grade. Water with a typical resistivity of 18 MΩ was produced from a Milli-Q purification system (Millipore, Les Ulis, France). Preparation of the Microporous Electrodes. The modified electrodes were prepared with very thin porous aluminum oxide films (a few micrometers) produced in the laboratory. Miller and Majda first described the procedure.10 In the present work the alkylation step was omitted. Briefly, aluminum oxide films were generated by anodic corrosion of aluminum foils in phosphoric acid (4% w/v) for 2 h at 65 V. The separation of the oxide film from the aluminum substrate and the removal of the barrier layer were performed according to Marchal et al.11 After the surface was rinsed in water and dried, amino groups were created at the surface of aluminum oxide by reaction with a freshly prepared ADMS solution in toluene (2%, v/v) for 8 h, and the surface was rinsed extensively with toluene and dried. They were transferred into a vacuum deposition apparatus (Edwards model E306A) where they were coated with ca. 2 µm thick gold films. The geometrical characterization of these films (thickness, pore density and pore size) was routinely performed by environmental scanning electron microscopy (ESEM ref XL 30 from Philips). Finally, the gold-coated oxide films were mounted on the tip of a glass tube (3 mm in diameter), using conductive silver glue for electrical contact. Just before use, the remaining bare surface of gold at the bottom of the pores was treated for partial alkylation with an alkyl mercaptan (OM). This treatment helps fusion of vesicles and decreases substantially the microporous electrode background current.11b To avoid total blocking of the quinone electrochemistry, the electrodes were dipped only for 2 min in a 1 mM solution of OM in ethanol/water (4/1, v/v) and rinsed with methanol and toluene to remove all unbound OM molecules before being finally dipped in phosphate buffer. Supported Bilayer Assemblies on the Streptavidin Sublayer. The self-assembly of the streptavidin sublayer on the (14) Gough, D. A.; Leypoldt, J. K. Anal. Chem. 1979, 51, 439-447.

Langmuir, Vol. 18, No. 8, 2002 3265 porous surface covered with amino groups was achieved in two steps. First the microporous electrode was dipped for 30 min in a 2 mM NHS-lc-biotin solution in a phosphate buffer (50 mM, pH 8). After extensive rinsing, the electrode was dipped for 10 min in a 0.1 µg/cm3 streptavidin solution in the PBS buffer (phosphate buffer 0.01 M, pH 7.4 + NaCl 0.15 M). After rinsing in a 50 mM OG solution and in the convenient buffer in three different baths, the electrodes were ready for experiments aimed at the fusion of vesicles. Mixed phospholipid-Q10 vesicles were obtained from dried lipids as follows: The chloroform solution of the different lipids, biotinylated DPPE and Q10 in the required ratios, was evaporated under nitrogen flow and dried under vacuum for 1 h at least. The film was resuspended from the walls of a glass tube by vigorous vortexing in 5 mL of the buffer. This solution was sonicated to clarity, four times for 3 min each, with a Branson model 250 sonicator (Danbury, CT) set at 60 W power, the temperature being maintained between 40 and 50 °C with a cold bath in case of need, with care being taken not to allow the temperature of the vesicles to drop below the gel-liquid-crystalline-phase transition temperature. The solution of biotinylated small unilamellar vesicles (SUVs) was cleaned of titanium particles by centrifugation at 3000g. The size of the vesicles was routinely measured by quasielastic light scattering with a Zetasizer 1000/3000 from Malvern Instruments (Malvern, U.K.). Provided that this measurement was performed less than 30 min after sonication, the average diameter of the SUV was regularly in the 30-40 nm range. After 4 h of storage, slow spontaneous vesicle fusion increased the average diameter to about 70 nm. To avoid this aging, the vesicle solutions were prepared just before use. For the formation of the supported bilayer, the microporous electrodes were transferred and incubated at the convenient temperature in a solution of 0.5-10 mM biotinylated vesicles for adsorption on the streptavidin sublayer. When performed, the PEG treatment consisted of 5-10 min of dipping of the rinsed electrode in a PEG 8000 solution (30% w/v in the buffer). It is worth emphasizing that repeated dipping of the electrode in the various solutions did not wash out the lipid layer; the pores are always filled with the solution when the tip of the electrode crosses the air/water interface. Surface Plasmon Resonance Analysis. The SPR instrument was a Biacore X (Biacore, Sweden) used with F1 sensor chips (Biacore) of flat gold surface. The sensor chips were pretreated with the 20 mM aminoethanethiol solution in water for 4 h. After the chips were rinsed by a 15 min injection of a 50 mM OG solution at a flow rate of 5 µL/min, the different solutions of reactants were introduced at a flow rate of 2 µL/min. Electrochemical Measurements. The anaerobic electrochemical cell contained three electrodes: the working microporous electrode, a platinum foil auxiliary electrode, and a KCl saturated aqueous calomel reference electrode (SCE ) 0.234 V vs the normal hydrogen electrode at 30 °C) to which all potentials are referred. Gentle bubbling of argon or nitrogen reduced the partial pressure of oxygen in the main compartment. The temperature was controlled at 30 °C by water circulation in the outer compartment of the water-jacketed cell. For the RDE experiments, the microporous electrodes were introduced in a special holder at the tip of the rotating device (motor and speed control unit CTV101T from Tacussel Radiometer Analytical, Villeurbanne, France). A PAR model 273 potentiostat controlled by a PC computer and a Power-Suite software package (Princeton Applied Research, Oak Ridge, TN) or a Voltalab 32 controlled by a PC computer and a Voltamaster software package (Tacussel, Radiometer Analytical,) was used for cyclic voltammetry, chronocoulometry, and RDE amperometry.

Results and Discussion Geometrical Characterization of the Porous Aluminum Oxide Films. Environmental scanning electron microscopy (ESEM) measurements were directly performed on the oxide samples without metal coating of the alumina surface. The structure, shown in Figure 1 as a regular array of parallel and roughly cylindrical pores

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Table 1. Geometrical Characterization of the Microporous Oxide pore densitya average pore diametera d, oxide thicknessa Fg, geometrical porosityb total surface oxide areab per cm2 of apparent area per cm3 of oxide volume per electrode of 0.07 cm2 a

(5.1 ( 0.5) ×109 pores per cm2 110 ( 10 nm 4.2 ( 0.4 µm 49 ( 8% 17 ( 2 cm2 per µm of thickness (1.7 ( 0.2) × 104 cm2/cm3 4.9 ( 0.7 cm2 (thickness 4.2 µm)

From ESEM images. b Calculated.

normal to the oxide surface, was characterized by three parameters: the pore density, the average pore diameter, and d, the oxide thickness (Table 1). A relative standard deviation of 10% was found for the measured parameters, either between oxide samples from a same batch or between samples from different batches. The geometrical porosity defined as the relative surface area of holes (Fg in percent) is derived from the pore density and the pore diameter. The total oxide surface area is calculated by adding all the inner surface areas of the cylindrical pores. It was previously demonstrated by radioactive labeling11a that the effective surface area of alkylated porous oxides is identical to the calculated surface area within the experimental uncertainties. Formation of the Streptavidin Sublayer. A threestep layer-by-layer assembly of the individual functional elements was applied to build the streptavidin sublayer starting from the inner pore surface of aluminum oxide. The three successive steps were (i) grafting the aminosilane (ADMS) on aluminum oxide, (ii) biotinylation of the surface amino group with NHS-lc-biotin, and (iii) docking of streptavidin by exposure to a diluted solution of streptavidin. Numerous optimizations of avidin/biotin constructions on solid supports have been reported.2a,15 Particularly the kinetics of protein loading on a flat gold surface can be followed by surface plasmon resonance (SPR).16 We used a similar approach to monitor the construction of the streptavidin sublayer on a flat gold surface in parallel to its construction on the inner walls of the pores. The corresponding experiments were carried out with a Biacore apparatus on gold chips bringing into play the same solutions and coating procedures as those used for the pores of aluminum oxide except that the first amine layer was grafted with aminoethanethiol (Figure 3A). The objective was to immobilize a discontinuous streptavidin sublayer in such a way that a small reservoir of aqueous phase could exist between the bilayer and the solid. The parameters were optimized so as to obtain streptavidin coverage of 10-15% of the maximum possible surface coverage. According to the calibration of Stenberg et al.16a or to the calculations of Sigal et al.,16b a thickness of 1.4 nm of protein (refractive index of 1.45) or a protein coverage of 100 ng cm-2 corresponds to 1000 RU (arbitrary units) of the Biacore apparatus meaning that 10-15% of the maximum surface streptavidin coverage should give between 350 and 550 RU. Therefore the SPR signal of 500 RU found in Figure 3A lies within the expected interval. (15) (a) Frey, B. L.; Jordan, C. E.; Kornguth, S.; Corn, R. M. Anal. Chem. 1995, 67, 4452-4457. (b) Hoshi, T.; Anzai, J.; Osa, T. Anal. Chem. 1995, 67, 770-774. (c) Bieri, C.; Ernst, O. P.; Heyse, S.; Hofmann, K. P.; Vogel, H. Nat. Biotechnol. 1999, 17, 1105-1108. (d) Anicet, N.; Bourdillon, C.; Moiroux, J.; Save´ant, J.-M. J. Phys. Chem. B 1998, 102, 9844-9849. (e) Fisher, M. I.; Tjarnhage, T. Biosens. Bioelectron. 2000, 15, 463-471. (16) (a) Stenberg, E.; Persson, B.; Roos, H.; Urbaniczky, C. J. Colloid Interface Sci. 1991, 143, 513-526. (b) Sigal, G. B.; Bamdad, C.; Barberis, A.; Strominger, J.; Whitesides, G. M. Anal. Chem. 1996, 68, 490-497.

Figure 3. Monitoring of the structure formation: experimental traces of the SPR responses given by the Biacore apparatus at 30 °C. (A) Successive surface binding of the NHS-lc-biotin arm and streptavidin on a gold surface modified with aminoethanethiol: arrow 1, introduction of a 2.1 mM NHS-lc-biotin solution in phosphate buffer; arrow 2, introduction of a 0.5 µg cm-3 streptavidin solution in PBS buffer; arrows OG, rinsing phases with a 50 mM octylglucoside solution in a convenient buffer made of 50 mM, pH 8 phosphate buffer at times e2200 s; 0.01 M, pH 7.4 phosphate buffer + 0.15 M NaCl (PBS buffer) in the 2200-4200 s time interval. (B) Continuation of the trace obtained during the loading of the streptavidin sublayer produced in part A with biotinylated vesicles: arrow 3, introduction of a 1 mM solution of eggPC/DMPE vesicles in 50 mM, pH 7, Tris-HCl buffer + 0.1 M NaCl. The vesicle composition is: 64.5% eggPC; 35% DMPE and 0.5% biotinylated DPPE in weights. The rinsing phases with a 50 mM OG solution in the same buffer wipe out the lipid structures, as expected.

Similarly, an estimation of the biotin coverage can be deduced from the measurement of 550 RU after NHSlc-biotin treatment. A simple calculation gives 1000-1500 RU for full coverage with NHS-lc-biotin, this evaluation being less reliable than that for a protein since the packing of the immobilized molecules is unknown in that case. However the rough estimate ascertains that the surface concentration of biotin is in large excess and the parameter allowing easy control of the streptavidin coverage is the low streptavidin concentration presented to the biotinylated surfaces. Blanks, omitting the NHS-lc-biotin step, demonstrate that nonspecific binding of streptavidin is negligible (less than 50 RU, not shown in Figure 3A) as expected with streptavidin at low concentration. Immobilization of Vesicles on the Streptavidin Sublayer. Immobilization, and possibly spontaneous fusion, of the biotinylated vesicles on the streptavidin sublayer can be performed. The loading of vesicles was preliminary explored by SPR measurements on a plane surface. Immobilization of lipidic material can thus be easily observed as shown in Figure 3B. We measured repetitively a response lying between 3500 and 4000 RU during sublayer exposure to solutions of vesicles at various


concentrations (0.5-10 mM in lipids containing 0.5 mol % of biotinylated lipids) during 5-20 min. The experimental data seem in accordance with the expected response for an adsorbed unilamellar bilayer, which has been found to give rise to 4000 ( 500 RU responses.17 However, in our opinion, this is more a coincidence than a definitive proof of the fusion of vesicles. According to the principle of SPR spectroscopy, the method gives only an overall view of the amount of lipids retained on the sublayer and cannot discriminate between anchored intact vesicles and connected bilayers. This is why we had to use electrochemical methods of characterization. Simultaneously we tested nonspecific binding (NSB) of vesicles on the streptavidin sublayer in numerous blanks performed with nonbiotinylated vesicles (not shown in Figure 3B). The results were found reliable only at low vesicle concentrations (up to 1 mM in lipids). The SPR responses were then less than 400 RU after 1 h of loading. This was expected for vesicles not containing biotinylated lipids, which have no specific affinity for the sublayer. But when the lipid concentrations were higher (5-10 mM), nonreproducible loadings and NSB responses at 5000 RU, or even more, revealed spontaneous adsorption of uncontrolled lipidic structures, possibly multilamellar. Electrochemical Monitoring of the Bilayer Formation. The progress of the fusion of vesicles and the formation of a continuous bilayer along the walls of the pores can be followed in real time by use of the microporous electrode. Ubiquinone (coenzyme Q10), a strictly waterinsoluble electrochemically active compound is dissolved at low concentration into the lipid leaflets of the SUV used for attachment to the streptavidin sublayer. The quantitative analysis of the electrochemical behavior of Q10 in similar structures has already been reported.18 At sufficiently low Q10 concentration, less than 2 mol % for the Q10/lipid ratio, it is well admitted that the Q10 molecules only lie and move within the hydrophobic mid-plane of the bilayer.11,19 The very long isoprenic chain (50 carbon atoms in the case of Q10) attached to the quinone forbids any motion and electrochemical reaction in solution. Thus the amount of Q10 detected electrochemically at the gold electrode only reflects the extent of fusion of the vesicles on the support (Figure 2). Moreover, the continuity of the supported bilayer can be ascertained by the measurement of the lateral diffusion coefficient of Q10. The spontaneous fusion of biotinylated DPMC/Q10 vesicles was monitored by means of cyclic voltammetry as a function of time at different temperatures. After selfassembly of the streptavidin sublayer on aluminum oxide, the microporous electrodes were loaded by dipping in the solution of vesicles for several hours. The amount of electrochemically addressable quinones was regularly measured in the presence of phosphate buffer in a separate cell (Figure 4A). The electrochemical reduction of the quinone is a complex process involving two electrons and two protons transfers, which has been analyzed previously in detail under similar conditions, and the nonreversible behavior of the hydroquinone/quinone redox couple embedded in the bilayer observed presently is quite similar to previous results obtained at the same pH.18b The (17) (a) Cooper, M. A.; Try, A. C.; Carroll, J.; Ellar, D. J.; Williams, D. H. Biochim. Biophys. Acta 1998, 1373, 101-111. (b) Radler, U.; Mack, J.; Persike, N.; Jung, G.; Tampe´, R. Biophys. J. 2000, 79, 3144-3152. (c) Keller, C. A.; Glasmastar, K.; Zhdanov, V. P.; Kasemo, B. Phys. Rev. Lett. 2000, 84, 5443-5446. (18) (a) Moncelli, M. R.; Becucci, L.; Nelson, A.; Guidelli, R. Biophys. J. 1996, 70, 2716-2721. (b) Marchal, D.; Boireau, W.; Laval, J.-M.; Moiroux, J.; Bourdillon, C. J. Electroanal. Chem. 1998, 451, 139-144. (19) Rajarathnam, K.; Hochman, J.; Schindler, M.; Ferguson-Miller, S. Biochemistry 1989, 28, 3168-3176.

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Figure 4. Electrochemical monitoring of supported bilayer formation by loading of the streptavidin sublayer with DMPC vesicles and spontaneous fusion. The loading is performed in the vesicles solution; the electrochemical measurements are carried out in a separated cell containing phosphate buffer. (A) Voltammograms at low potential scan rate (5 mV s-1) of Q10 solubilized in the mid-plane of the bilayer after different times of loading. Experimental traces at 30 °C after 1, 72, 96, and 144 h for curves a, b, c, and d, respectively. The composition of the mixed SUV vesicles used for the loading is 97.5% DMPC, 2% Q10, 0.5% biotinylated DPPE in weights. They are introduced at 1 mM concentration in 0.1 M, pH 7, phosphate buffer. Integration of the cathodic peaks gives Q10 electric charges of 0.5, 1.4, 2.0, and 4.0 µC for curves a-d, respectively. Dotted line: background current in phosphate buffer before loading. Note that despite the OM treatment, the anodic background current increases after long-term use. (B) Effect of temperature and detergent on the kinetics of lateral fusion monitored by measurement of Q10 electric charges. Temperatures are 30 °C (9) and 50 °C (b, O). The detergent is 1 mM OG (b) at 50 °C. The broken line at 8.9 µC shows the theoretical ubiquinone loading expected for complete lateral fusion.

potential scan rate used in cyclic voltammetry is sufficiently slow to ascertain that the area under the cathodic peak gives the total amount of Q10 able to reach the electrode by lateral diffusion from the whole microporous structure. As shown in Figure 4, spontaneous fusion, which ensures progressive connection between the quinone pool and the gold surface, requires a long time (several days). Its kinetics is markedly different from the kinetics of im-

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mobilization of vesicles followed by SPR (Figure 3B), in the latter case the time scale is much shorter (tens of minutes). Considering the formation of a supported membrane on hydrophilic surfaces by random deposition of vesicles present in solution, Radler et al.20 proposed a four-step mechanism: (a) vesicle approach; (b) adhesion; (c) vesicle rupture; (d) lateral spreading and membrane fusion. In our case the cyclic voltammetry and SPR techniques enabled us to follow the progress of different steps of the overall process. The SPR results indicate that the capture of biotinylated vesicles by the streptavidin sublayer including the first two steps of the preceding process, approach and adhesion, is completed within a relatively short time and is not rate limiting. On the other hand, cyclic voltammetry shows that expansion of the supported bilayer creating a lateral connection with the bottom of the pores is 2 orders of magnitude slower, thus establishing that the rupture of the vesicles and/or the lateral spreading of the bilayer are or is rate limiting. The structural reorganization of the attached lipid material does not cause apparent mass change as detected in SPR spectroscopy. A similar interpretation has been already proposed to justify the kinetics of a thiolipid attached bilayer formation, which was monitored in parallel by impedance measurements and SPR spectroscopy.21 The results reported in Figure 4 thus confirm the four-step mechanism mentioned above and point out the ratelimiting steps. That at least one of the last two steps is rate limiting is further assessed by the following results. The rate of lateral connection in the attached structure is significantly increased when the vesicles are slightly destabilized, either by a temperature increase from 30 to 50 °C or by addition of a low concentration of detergent in the buffer (1 mM OG, i.e., far below the OG critical micellar concentration). Moreover, fusion of vesicles is negligible (not shown in Figure 4) at 15 or 20 °C, temperatures that are lower than the phase transition temperature of DMPC (24 °C). It is worth mentioning that a fast lipid/quinone loading is systematically measured at the beginning of the experiments (see for example cyclic voltammogram a in Figure 4A). The corresponding preliminary fusion takes place probably on the alkylated gold surface located at the bottom of the pore. The high efficiency of the fusion of vesicles on hydrophobic surfaces such as OM-modified gold is well documented,3c,22 and it may trigger bilayer formation from the bottom of the pore. That is one of the reasons why we modified the gold surface at the molecular level as indicated by the OM layer representation given in Figure 1C. However the most striking result is that we never achieved full lipid/quinone coverage by spontaneous fusion of vesicles on the streptavidin sublayer whatever the duration we were able to allow experimentally to the process. The maximum theoretical coverage can be calculated once the total area of the walls of the pores (4.7 cm2), the quinone-to-lipid ratio (2.0 mol %), and the mean area occupied by a lipid in a bilayer (about 67 Å2)4a,11 are known. That gives a Q10 surface concentration at maximum theoretical coverage of 9.8 pmol cm-2 and an area under the cathodic peak in cyclic voltammetry equal to an electric charge of 8.9 µC. Despite uncertainties of ca. 20% on the parameters used in the preceding calculation, the (20) (a) Radler, J.; Strey, H.; Sackmann, E. Langmuir 1995, 11, 45394548. (b) Feder, T. J.; Weissmuller, G.; Zeks, B.; Sackmann, E. Phys. Rev. E 1995, 51, 3427-3433. (21) Lang, H.; Duschl, C.; Vogel, H. Langmuir 1994, 10, 197-210. (22) Kalb, E.; Frey, S.; Tamm, L. K. Biophys. Biochim. Acta 1992, 1103, 307-316.


saturating value of 5-6 µC found experimentally (Figure 4B) is significantly smaller than that the one corresponding to full coverage. The simplest explanation could be that the limitation results from incomplete coverage or more precisely from the coexistence of connected areas with areas occupied by nonruptured vesicles as already observed by Radler et al.20a Another explanation could be the occurrence of long-term addition to the supported lipid bilayer of single DMPC molecules existing in solution. As a result the Q10/lipid ratio within the supported bilayer would decrease. Whatever the reason, the spontaneous formation of the supported DMPC bilayer is clearly too slow for useful incorporation of active intrinsic proteins from proteoliposomes. In consequence, we explored two alternative approaches. In the first one, not successful, we tried to apply numerous variants of classical vesicle reconstitution procedures that end up with removal of the detergents.23 Briefly, the microporous electrodes were dipped in a solution of lipids (typically, 10-3 M DMPC, 2 × 10-5 M Q10, 5 × 10-6 M biotinylated DPPC) solubilized in a detergent at a high concentration (50 mM OG in phosphate buffer). The electrodes still dipped in the solution; the excess of detergent was then eliminated either by dialysis or by adsorption on Biobeads. Electrochemical measurements gave disappointing electric charges of 0.5-1.5 µC. We gave up after numerous unsuccessful attempts involving various lipid/detergent ratios and different durations for detergent removal, from an hour to 1 day. It turned out that the second approach, which is described in the following as the method of triggered fusion of vesicles, proved successful. Formation of the Supported Bilayer by Triggered Fusion of Vesicles. Our results show that the inefficiency of the spontaneous fusion of biotinylated vesicles on a streptavidin sublayer is due to the extreme slowness of fusion of the avidin/biotin attached vesicles. Incidentally, it is worth mentioning that this kind of attachment has been proposed for the immobilization of intact vesicles on various surfaces and polymers.24 With SPR measurements proving that the lipid material is promptly in place in the configuration of attached vesicles, we must find a way to trigger the fusion between the immobilized lipid material in a distinct second step, as indicated in Figure 2. Triggered fusion between vesicles in solution can be considered as a model for the elucidation of processes of biological fusion and is well documented.13 In particular, PEG-mediated fusion of model membranes has been shown to mimic the biomembrane process.25 From the mechanistic point of view, PEG is considered to force vesicles together due to depletion attraction; the vesicles are flattened by osmotic withdrawal of water, and finally fusion occurs at the highly curved boundaries between closed bilayers.13,25 The related studies provided evidence of the crucial influence of the lipid composition on PEGmediated fusion of SUVs, for example, the presence of phosphatidylethanolamine heads was shown to contribute greatly and we adapted accordingly the lipid composition of the vesicles prepared in the present work. We proceeded as follows. The streptavidin sublayer was first loaded 1 h with biotinylated eggPC/DMPE vesicles (23) Paternostre, M. T.; Roux, M.; Rigaud, J. M. Biochemistry 1988, 27, 2668-2677. (24) (a) Jung, L. S.; Shumaker-Parry, J. S.; Campbell, C. T.; Yee, S. S.; Gelb, M. H. J. Am. Chem. Soc. 2000, 122, 4177-4184. (b) Percot, A.; Zhu, X. X.; Lafleur, M. Bioconjugate Chem. 2000, 11, 674-678. (25) (a) Lee, J.; Lentz, B. R. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 9274-9279. (b) Haque, M. E.; McIntosh, T. J.; Lentz, B.R. Biochemistry 2001, 40, 4340-4348.


Langmuir, Vol. 18, No. 8, 2002 3269

Figure 6. Geometrical models for the decreases in apparent porosities that accompany the achievement of various steps in the assembly process. Schematic front view of a pore at the end of step 1, which consists of loading of the streptavidin sublayer with vesicles, and at the end of step 2, which consists of PEG mediated fusion of the immobilized vesicles. The representation reflects roughly the relative sizes of all the individual components involved in the construction.

Figure 5. Electrochemical monitoring of Q10 loading at different steps of the PEG-triggered supported bilayer formation from eggPC/DMPE vesicles. The loading was performed in the vesicles solution; the electrochemical measurements were carried out in a separated cell containing phosphate buffer at 30 °C. (A) Voltammograms at 5 mV s-1 after 1 h of dipping of the microporous electrode in a 5 mM solution of mixed lipid vesicles prepared in phosphate buffer (63.5% eggPC, 34% DMPE, 2% Q10, 0.5% biotinylated DPPE in weight): curve a, after rinsing in the buffer and before PEG treatment (ubiquinone electric charge deduced from the cathodic peak ) 1.7 µC); curve b, after PEG treatment consisting of a 5 min dipping in a solution of PEG 8000 at 30% w/v in buffer and rinsing (ubiquinone electric charge deduced from the cathodic peak ) 9.4 µC). (B) Fusogen effect of the PEG treatment: (b) loading and treatment as in part A; (O) blank using the same solution of eggPC/DMPE vesicles but without PEG treatment; (9) blank with 1 h PEG treatment but without introduction of DMPE into the vesicles. The broken line at 8.9 µC indicates the expected theoretical ubiquinone loading expected for complete lateral fusion.

by introduction of the microporous electrode into a solution of such vesicles and rinsing. The fusion was then triggered by dipping in a (30% w/v) PEG solution for 5 min followed by rinsing. Evidence of the efficiency of the PEG-mediated fusion is provided in Figure 5. After the 5 min triggering treatment, the Q10 related electric charge of 9.1 µC reaches the level corresponding to maximum theoretical coverage. In a set of control experiments, we checked out that the Q10 charge is, as expected, proportional to the Q10 to lipid ratio in the vesicles in the 1-3 mol % range (not shown

in Figure 5). Other controls confirmed that the fusion of attached vesicles follows the same rules as those applying to the fusion of vesicles in solution, namely: (i) pure DPMC vesicles are not sensitive to PEG mediated fusion whatever the temperature, above or below the transition temperature of 24 °C; (ii) the PEG treatment does not trigger the fusion of eggPC vesicles in the absence of DMPE (Figure 5B). Last, but not least, the lateral fluidity of the supported bilayer can be assessed by chronocoulometric measurement of the lateral diffusion coefficient of Q10. The method, which was previously described in detail in the case of supported hemi-bilayers,11b gives access to the diffusion coefficient (DQ) as does fluorescence recovery after the photobleaching technique (FRAP) with the important difference that labeling of the ubiquinone with a hydrophilic fluorophore may alter significantly the configuration of the ubiquinone molecule within the bilayer in the FRAP technique.11b With four distinct microporous electrodes, we found DQ ) (3 ( 1) × 10-8 cm2 s-1 at 30 °C, a value falling within the range of long-range diffusion coefficients measured by FRAP in vesicles going from 1.1 × 10-8 cm2 s-1 for a fluorescent derivative of Q1019 to 5.5 × 10-8 cm2 s-1 for a fluorescent derivative of Q2.1 Indirect Geometrical Characterization of a Pore by Measurement of the Electrochemical Porosity. It is expected that the size of the tethered assembly may not be negligible compared to the 110 nm average diameter of the cylindrical pores of aluminum oxide. The bilayer itself is 5 nm in thickness, streptavidin can be taken as a globule of ca. 5 nm diameter,26 and the biotinylated “longarm” spacer, which is long enough to allow unhindered interaction between its biotin head and avidin or streptavidin, is 2.3 nm in length.27 If all components are piled up from the pore wall, as sketched in Figure 6, the inner pore diameter could be decreased by (2.3 + 5 + 2.3 + 5) × 2 = 30 nm. Such a decrease in the average diameter of the pores through which the electrode and the bulk solution communicate can be detected experimentally by electro(26) Darst, S. A.; Ahlers, M.; Meller, P. H.; Kubalek, E. W.; Blakenbourg, R.; Ribi, H. O.; Ringsdorf, H.; Kornberg, R. D. Biophys. J. 1991, 59, 387-396. (27) In Bioconjugate techniques; Hermanson, G. T., Ed.; Academic Press: New York, 1996; pp 371-400.

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chemical measurement of the diffusion flux of a small water-soluble electrochemically active molecule. It has been shown that a Koutecky-Levich analysis of RDE data can be used for the determination of porosity in similar configurations.10a At an appropriately chosen electrode potential, the method is valid and accurate only when it is possible to reach electrode rotation rates that are high enough to create a diffusion layer so thin that the recorded stationary current is controlled by the rate of diffusion of the electrochemically active molecule through the pores. We chose ruthenium hexamine, Ru(NH3)63+, as the sensing molecule introduced in solution for the following two reasons. First the rate of heterogeneous electron transfer between the soluble Ru(NH3)62+/Ru(NH3)63+ redox couple and the gold electrode is rather fast. Second, both the reduced and oxidized forms of the redox couple bear electric charges and are very hydrophilic molecules; consequently they are not likely to undergo partition between the aqueous solution and the lipid bilayer. The formal potential of the Ru(NH3)62+/Ru(NH3)63+ redox couple is 0.180 V/SCE. When the electrode potential is negative enough to give a cathodic current that is potential independent, each Ru(NH3)63+ molecule reaching the gold electrode surface is reduced at this location and mass transfer proceeds at a steady state. Then10a,14

d 1 1 ) + ilim FACDFe 0.62FACD2/3ν-1/6ω1/2

(1)

ilim being the cathodic plateau current (in A), d the thickness of the aluminum oxide (in cm), A the electrode geometrical surface area (0.07 cm2), C the bulk concentration of Ru(NH3)63+ (in mol cm-3), D the diffusion coefficient of Ru(NH3)63+ (in cm2 s-1), Fe the electrochemical porosity equal (in %) to the sum of the effective areas of the cross sections of the cylindrical pores over the geometrical electrode surface area which supports them, ν the viscosity of the buffer solution at 30 °C (0.980 × 10-2 P), and ω the angular rate of the rotating electrode (in rad s-1). The diffusion coefficient of Ru(NH3)63+ in Tris/HCl buffer was carefully determined at a bare gold disk electrode of known geometrical area. We found (7.0 ( 0.4) × 10-6 cm2 s-1 at 20 °C and (8.1 ( 0.4) × 10-6 cm2 s-1 at 30 °C in good agreement with the literature (7.1 × 10-6 cm2 s-1 in 0.1 M KCl at 20 °C).28 A Tris/HCl (+ NaCl, 0.1 M) pH 7 buffer was used instead of phosphate (+ NaCl, 0.1 M) pH 7 buffer, the temperature dependence of the Ru(NH3)63+ diffusion coefficient appearing complex in the phosphate buffer, probably due to ion-pairing formation. Application of eq 1 indicates that Fe can be deduced from the extrapolated origin intercept of the 1/ilim versus 1/ω1/2 Koutecky-Levich plot. Koutecky-Levich plots revealing the porosity changes that accompany various steps in the process of loading or unloading the selfassembly are given in Figure 7. Typically, Fe decreases from 55% to 51% after the NHS-lc-biotin + streptavidin treatment and then goes down to 31% after the vesicle loading and 24% after the PEG treatment. The porosity of 51% found in the presence of the sole streptavidin sublayer is restored after destruction of the bilayer by rinsing with a detergent like OG at 50 mM. With the relative uncertainty on the thickness of an individual oxide being 10% at least, the relative uncertainty on Fe reaches (28) (a) Licht, S.; Cammarata, V.; Wrighton, M. S. J. Phys. Chem. 1990, 94, 6133-6140. (b) Mirkin, M. V.; Arca, M.; Bard, A. J. J. Phys. Chem. 1993, 97, 10790-10795.

Figure 7. Typical Koutecky-Levich plots allowing the measurement of the apparent porosity of a given microporous electrode at different phases of the supported bilayer assembly. Plots are obtained with (a) untreated microporous electrode, Fe ) 55%, (b) loaded with the streptavidin sublayer, Fe ) 52%, (c) after 1 h in the biotinylated vesicle solution, Fe ) 31%, (d) after PEG treatment, Fe ) 24%, and (e) after detergent rinsing, Fe ) 51%. The limiting currents were measured at -0.65 V/SCE with a 4.14 mM Ru(NH3)63+ solution in pH 7 Tris/HCl buffer at 30 °C. The rotation speeds were varied from 500 to 3000 rpm. The dashed line shows the data obtained at a bare gold disk electrode in an experiment that was carried out in order to determine the diffusion coefficient of Ru(NH3)63+ under the same conditions.

15% when going from one microporous electrode to another. However the relative uncertainty falls down to 5% for each set of measurements concerning a given oxide, and the relative changes observed at each step of the selfassembly process are remarkably similar when going from one microporous electrode to another. We took great care to ascertain that the experimental conditions required for correct use of eq 1 were fulfilled. Particularly, the electrochemical reduction of Ru(NH3)63+ must be fast compared to its diffusion within the pores. As the gold surface is modified by the OM treatment and possible bilayer connection at the bottom of the pore, there is a risk of severe decrease in the rate of heterogeneous electron transfer between the Ru(NH3)62+/Ru(NH3)63+ redox couple and the electrode at the modified gold surface. We proceeded to two types of verifications. First, the applied potential was systematically varied to ensure that a plateau in the measured porosity was reached. Second, control experiments were carried out at OM-untreated microporous electrodes cleared of their lipid coating on gold by anodic stripping of the gold surface at 0.5 V/SCE at each stages of the bilayer assembly. The corresponding results are summarized in Figure 8 and Table 2. A porosity plateau is easily reached at microporous structures not bearing the supported bilayer upon decreasing the electrode potential whatever the treatment applied to the gold surface. The light OM treatment does not alter the electrochemical behavior of the Ru(NH3)62+/ Ru(NH3)63+ couple; the cyclic voltammograms (not shown) at bare and OM treated microporous electrodes are identical. After the last step of the assembly process (after PEG in Figure 8), it is not obvious that a plateau value


Langmuir, Vol. 18, No. 8, 2002 3271

Table 2. Comparison between Geometrical and Electrochemically Measured Apparent Porosities, Microporous Electrodes at Different Stages of the Loading Process

geometrical porosity bare microporous electrode + streptavidin sublayer + biotinylated vesicles + PEG treatment (supported bilayer) + detergent

(Fg)a

49 ( 8% (0) 47% (1 nm in av) no model 27% (2 + 5 + 2 + 5 nm) 47% (1 nm in av)

measured porosity (Fe)b

Fe measured in the absence of OMc

55 ( 9% 51 ( 9% 31 ( 15% 24 ( 4% 50 ( 9%

54 ( 9% 52 ( 9% 40 ( 10% 26 ( 4% 51 ( 9%

a Deduced from Table 1 and Figure 6. The apparent thickness used for each layer are given in parentheses (see text). b From the electrochemical determination (average for eight electrodes). c From the electrochemical determination with OM-untreated gold and after electrochemical cleaning (average for three electrodes).

ible when going from one microporous electrode to another than the porosity measured after final PEG treatment. That brings further confirmation that the PEG treatment improves the self-organization of the final lipidic structures, among other possible effects, and that the final characteristics of the supported bilayer do not depend significantly on those of the structures created in the intermediate phases of the loading. Finally, we can calculate the minimal amount of lipids (or vesicles) needed to produce a continuous bilayer in the template at the end of the process: According to Table 1, the inner surface of one pore is π × (110 × 10-7) × (4.2 × 10-4) ) 1.45 × 10-8 cm2. The surface of one vesicle is π × (40 × 10-7)2 ) 5 × 10-11 cm2. One pore surface is thus equivalent to almost 300 vesicles of diameter 40 nm. The fact that PEG provokes the formation of a continuous bilayer is convincing proof that the pores are regularly filled with very many vesicles before PEG treatment. This confirms our view of bilayer formation (Figures 2 and 6). Conclusion

Figure 8. Apparent porosities (Fe) deduced from the KouteckyLevich plots. Same experimental conditions as in Figure 7. The measurements were carried out with two microporous electrodes made from the same batch of oxide production: (b) electrode normally treated with octadecyl mercaptan (OM); (O) the OM treatment was omitted and an electrochemical cleaning was performed before each measurement to get rid of possible blocking of the gold surface by lipid structures.

is obtained for the measured porosity of the microporous electrode bearing the expected molecular construction, including the supported bilayer. It is quite likely that the bilayer covers the gold surface as depicted in Figures 1 and 6. However, electrochemical cleaning of the gold surface enables us to ascertain that the porosity plateau is actually reached at the most negative potential as shown in Figure 8. The porosity then measured is Fe ) 26% (Table 2), in remarkably good agreement with the theoretical porosity (Fg) of 27% that can be calculated according to the geometrical model assuming that the bilayer (5 nm) is supported at a distance of about 2 + 5 + 2 nm from the pore inner wall. After vesicle loading and before PEG treatment, there is no realistic geometrical model which can be considered for the pores that are partially blocked by intact vesicles as sketched in Figure 6. For example we do not know if the anchored vesicles are still spherical or begin to flatten against the streptavidin sublayer. Consequently, the measured porosity cannot be related to a defined geometry of the space that can be filled by the solution inside the pores. Moreover, the porosity measured after vesicle loading and before PEG treatment is much less reproduc-

Spontaneous fusion and connection of lipid vesicles on a protein sublayer is a very slow process even when an additional driving force like biotin/avidin affinity is used to increase the local concentration of the lipid material. We propose a rapid method for successful assembly of tethered and supported lipid bilayers. The method involves two successive and easily controlled steps. Triggering of the fusion with a fusogen agent follows accumulation and immobilization of the vesicles on the template. As already reported in a preliminary note,12 the presence of a PEG fusogen in solution triggers very efficiently the connection between anchored vesicles in order to produce smooth supported bilayers. The quality and the continuity of the supported bilayer are assessed by means of three complementary types of controls. RPS measurements show that the attachment of vesicles containing biotinylated lipids to a flat surface of gold covered with a streptavidin sublayer proceeds at a relatively fast rate and that the amount of immobilized lipid material corresponds approximately to that required for the formation of a supported bilayer. When ubiquinone Q10 is initially introduced into the hydrophobic part of the vesicles, electrochemistry at microporous electrodes allows the determination of the amount of ubiquinone of electrochemically addressable Q10 and the rate of its long range lateral diffusion in the final structure. Both reflect the extent to which continuity is created within the supported bilayer. The electrochemical measurements reveal that complete fusion is achieved only after PEG triggering. The lateral diffusion coefficient of 3 × 10-8 cm2 s-1 at 30 °C is as expected for a model membrane. Finally the porosity of the microporous structure is also geometrically characterized before and after bilayer assembly. The

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electrochemical measurement of the diffusion fluxes of water-soluble ruthenium-hexamine through the pores gives the average inner diameter of the pores. As expected each step of the assembly process causes a decrease in the apparent inner diameter and the final porosity measured in the presence of the supported layer is found in good agreement with a geometrical model taking into account the known sizes of a smooth lipid bilayer, streptavidin, and the biotinylated spacers. Compared with a bilayer supported on flat surface, the compaction of the biomimetic structure can be increased by several orders of magnitude when the support is a microporous structure similar to the one contained in the microporous electrode we used. About 20000 cm2 of continuous supported bilayer can be included within 1 cm3 of aluminum oxide, a surface-to-volume ratio that is


only 1 order of magnitude smaller than in natural organelles such as mitochondria (2 × 105 cm2 cm-3). It is expected that the approach elaborated in the present work can help greatly in the development of new strategies concerning the reconstitution and the monitoring of functional membrane proteins such as the biological complexes involved in the electron-transfer chains. Acknowledgment. We acknowledge the Chemical Engineering Department of UTC for permanent access to the Zetasizer apparatus. The support of this research by the CNRS through PCV Grant 98-091 is gratefully acknowledged. LA011585T

Formation of Tethered and Streptavidin-Supported Lipid Bilayers on a

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