In Situ Investigations of the Formation of Mixed Supported Lipid

Consequently, the rate of bilayer formation of the 70/30 mixture is faster than the rate for the 50/50 mixture. A complete coverage is observed at t )...
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NANO LETTERS

In Situ Investigations of the Formation of Mixed Supported Lipid Bilayers Close to the Phase Transition Temperature

2004 Vol. 4, No. 1 5-10

B. Seantier,† C. Breffa,† O. Fe´lix,*,† and G. Decher*,†,‡ Centre National de la Recherche Scientifique (CNRS), Institut Charles Sadron, 6 rue Boussingault, F-67083 Strasbourg-Cedex, France, and UniVersite´ Louis Pasteur (ULP), 1 rue Blaise Pascal, F-67008 Strasbourg-Cedex, France Received August 1, 2003; Revised Manuscript Received November 14, 2003

ABSTRACT The formation of supported phospholipid bilayers (SPBs) has been studied by atomic force microscopy (AFM) and by dissipation-enhanced quartz crystal microbalance (QCM-D). AFM experiments on mixed bilayers of 1,2-dimyristoyl-sn-glycero-3-phosphatidylcholine (DMPC) and 1,2-dipalmitoyl-sn-glycero-3-phosphatidylcholine (DPPC) permit to visualize in situ the process of bilayer formation, starting with vesicle adsorption to the formation of bilayer patches and finally to complete bilayers. QCM-D experiments using the same systems show how the process and its kinetics are controlled by temperature.

The preparation of ultrathin organic films on solid supports is interesting with respect to fundamental research as well as for applied research in material and life science.1 For several years, we have been studying layer-by-layer adsorbed films.2 Such multilayer assemblies may be composed of polyelectrolytes, colloids, or lipid bilayers.3-6 The latter systems are of special biological interest as they are similar to cell membranes which play a fundamental role in the creation of compartmentalized environments and which are thus at the base of life as we know it. In general, the structure of such membranes is called the “fluid mosaic model”, which describes a liquid-crystalline lipid bilayer.7 The interest in understanding the physical and physiological properties of cell membranes has spurred intense research for building simple model systems mimicking biological membranes. Many kinds of model systems have been used, including liposomes or lipid vesicles, black lipid membranes, Langmuir-Blodgett layers and supported lipid bilayers, especially composed of phospholipids (SPBs).8,9 Each system provides advantages and disadvantages with respect to preparation, properties mimicked, or applicable characterization techniques. Vesicles have mostly been used to study membrane transport, and black lipid membranes have proven useful for carrying out electrical measurements on lipid bilayers. More recently, planar supported lipid bilayers have been introduced.9,10 These two-dimensional lipid membranes (thickness about 5-6 nm) are typically formed on silica surfaces either * Corresponding authors. E-mail: [email protected]; felix@ ics.u-strasbg.fr. † Institut Charles Sadron. ‡ Universite ´ Louis Pasteur. 10.1021/nl034590l CCC: $27.50 Published on Web 12/16/2003

© 2004 American Chemical Society

as free bilayers or as so-called tethered bilayers which feature a small percentage of lipids covalently connected to the surface. The bilayers are separated from the solid support by an ultrathin film of water (1-2 nm) embedded between the solid support and the lower leaflet of the bilayer. It acts as a lubricant allowing both leaflets to remain fluid.11 As a consequence, many properties (structural and dynamic) of free vesicles or cell membranes are retained in the supported bilayers. Another major advantage of such system is that they exist at the solid/solution interface, which permits the use of a large number of characterization techniques. These tools include surface plasmon resonance (SPR),12 quartz crystal microbalance (QCM),13 impedance spectroscopy14 attenuated total reflection Fourier transform infrared spectroscopy (ATR-FTIR),15 scanning probe microscopy,16,17 and fluorescence recovery after photobleaching (FRAP)18,19 The most common methods for the preparation of SPBs are the fusion of vesicles on a hydrophilic silica surface,10,16,20 on a hydrophobic self-assembled monolayer,13,21 or on a predeposited hydrophobic lipid monolayer that is prepared by the transfer of a Langmuir monolayer.22 Depending on several factors such as the surface, the lipid composition, and other experimental conditions, the sequence of events during SPB formation (adsorption, rupture, and spreading) can be interrupted at intermediate stages. Interest in how these processes take place and what driving forces are involved has increased during the past few years. Several hypotheses concerning the mechanism of SPB formation have been advanced.9,17,20,23 In recent years some clues have been given, especially due to the use of in situ AFM.

While cell membranes are composed of many different components (proteins, several lipids, cholesterol, etc.), many physicochemical studies have been carried out using singlecomponent lipid bilayers or ill-defined lipid mixtures such as EggPC. While work on the phase diagram of lipid mixtures exists for bilayers dispersed in bulk water or buffer,24-26 there are only few studies on structure, phase separation, and kinetics of formation of SPBs composed of defined lipid mixtures. In single-component systems, the phase transition temperature (Tm) is dependent on the chemical nature of the lipid, especially on the length, degree of saturation, and branching of the alkyl chain and on the chemical structure of the headgroup. In multicomponent systems, the phase diagram is strongly influenced by the composition and miscibility of lipid mixture.24,25,27 The mixing behavior of different components is reflected by the shape of the phase diagram of the mixture as typically obtained by differential scanning calorimetry (DSC). The phospholipids used in the experiment described here, 1,2dimyristoyl-sn-glycero-3-phosphatidylcholine (DMPC) and 1,2-dipalmitoyl-sn-glycero-3-phosphatidylcholine (DPPC), show an ideal mixing in bulk. The transition temperature as a function of lipid composition was investigated by Blume for the case of multilamellar bilayers.26 Transition temperatures, however, are expected to depend on vesicle size or on the substrate in the case of SPBs. Our experiments show, in agreement with previous reports,26,28 that the phase transitions of SUVs are broadened and shifted to slightly lower temperature. It is generally assumed that the formation of supported bilayers requires a temperature above the phase transition temperature of the chosen lipid or lipid mixture. While most of the studies on SPBs were carried out at temperature way above the Tm, we have chosen to work close to the Tm in order to control the kinetics of the processes involved. To our knowledge, there is only one study on SPB formation of a lipid mixture below its phase transition temperature.29 Here, we present for the first time, the direct visualization of SPB formation from unilamellar vesicles close to their phase transition temperature as studied by AFM and by dissipation-enhanced quartz crystal microbalance (QCM-D) measurements. These techniques, which are widely used analytical surface tools, give detailed complementary in situ information on vesicle adsorption, rupture, and bilayer formation. Due to technical limitations of both techniques, experiments were carried out on the chemically similar but not identical surfaces, mica (AFM) and amorphous SiO2 (QCM-D). However, both surfaces are known to permit the formation of supported bilayer membranes via a different kinetic pathway under similar conditions. On mica, SPB formation proceeds by smaller vesicles first fusing to larger ones before they rupture and form supported membranes.17 On the other hand, on SiO2, QCM-D30 and AFM31have suggested a pathway through a critical coverage of vesicles on the surface. The latter can also be strongly affected by addition of a fusogenic agent such as CaCl2 or MgCl2 into the buffer.31-33 6

The visualization of the formation of supported lipid bilayers using AFM was reported by Jass et al. in 2000.16 The slow kinetics of the fusion of unilamellar vesicles (200400 nm) on silicon wafers has permitted to propose a mechanism from vesicle adsorption to lipid bilayer patches via flattened liposomes. Different studies have shown that the rupture process is influenced by the vesicle diameter17 and the nature of the supporting surface.13,20,30 In our approach, SPBs were formed by rupture of small unilamellar vesicles (50 nm). Time-lapse in situ contact AFM imaging allows visualization of nanoscopic bilayer patches, which are important intermediates during the formation of a “complete” bilayer. These experiments on SPB formation were performed on DMPC/DPPC mixtures (ratios 70/30 and 50/50, respectively) in the vicinity of the phase transition temperature. For the experiments reported here, multilamellar vesicles (MLVs) were prepared to a final concentration of 2 mg/mL by hydration of the corresponding lipid film with buffer (Tris‚ HCl 10 mM, NaCl 150 mM, CaCl2 2 mM, adjusted to pH ) 7.4 by NaOH 0.5 M). The lipid film was obtained by mixing appropriate amounts of lipids dissolved in chloroform, evaporating the organic solvent in a nitrogen flow, and drying overnight under vacuum. The phase transition temperatures of the lipid mixtures were determined by differential scanning calorimetry (microDSC III, Setaram) on MLV suspensions of 2 mg/mL and were in agreement with the literature.24 For DMPC/DPPC MLV mixtures, the phase transition temperatures were determined to be 29 °C and 33 °C (peak center of heating run at 0.25 deg./min) for a composition of 70/30 and 50/50, respectively. Small unilamellar vesicles (SUVs) were produced from 2 mL MLV suspensions by sonication to clarity (12 min, continuous mode) at room temperature with a tip sonicator (Branson Sonifier B15 cell disrupter, Danbury, CT). Titanium particles were removed by centrifugation (10 min at 5000 rpm). The clear SUV suspension was diluted with buffer to a final concentration of 0.05 mg/mL that was used for all further experiments. The size of the formed SUV vesicles was measured at 25 °C by dynamic light scattering (Malvern Zeta Sizer 300HS) at a scattering angle of 90°. The average diameter obtained for all lipid mixtures and pure lipids was approximately 50 nm. A Digital Instruments (Santa Barbara, CA) Nanoscope III AFM fitted with a 10 µm scanner (E-scanner) and a contact mode liquid cell was used to image in situ SPB formation. Liquid solutions were exchanged using the conventional O-ring in order to avoid leakage. Standard Si3N4 tips with a nominal force constant of 0.12 N/m were used. Force and scanning speed (3 Hz with 512 × 512 data points) were optimized for minimum damage and maximum resolution. Images were recorded in both height and deflection mode. Height images were flattened and plane adjusted. All measurements were done on the height images. For the AFM studies, SPBs were formed at room temperature (24 °C), which is 5 and 9 degrees below the phase transition temperature of the respective mixtures. Freshly cleaved mica (Metafix, Montdidier, France) covered with buffer was used Nano Lett., Vol. 4, No. 1, 2004

Figure 1. Time-lapse series of AFM images of DMPC/DPPC 50/ 50 demonstrating SPB formation via intermediate nanoscopic patches (A ) 0 min, B ) 11 min, C ) 17 min, D ) 19 min, E ) 28 min, F ) 62 min). Because of a noncomplete surface coverage after 30 min, a second injection was needed to obtain the bilayer (1F). The cross section 1G identified by the line in 1B shows that the height of SPB patches is approximately 5 nm.

as reference surface (Figures 1A and 2A). Vesicle solutions were introduced into the liquid cell using a syringe and the formation of bilayers was followed by time-lapse AFM for 1 h. In all cases, the adsorption, fusion, and rupture process of vesicles leads to clearly visible SPB patches and allows the analysis of the bilayer thickness as well as the evolution of the patches size and shape in time. The height profile indicates two levels with a step height corresponding to a lipid bilayer. Nano Lett., Vol. 4, No. 1, 2004

Figure 2. Time-lapse series of AFM images of DMPC/DPPC 70/ 30 demonstrate a considerably faster SPB formation via intermediate nanoscopic patches (A ) 0 min, B ) 14 min, C ) 17 min, D ) 23 min, E ) 25 min, F ) 34 min) compared to 50/50 ratio (see Figure 1). The cross section 2G identified by the line in 2C shows that the height of SPB patches is approximately 5 nm.

Figure 1 shows an AFM image sequence for a mixture of DMPC/DPPC (50/50) during SPB formation. Round patches of similar size are present on mica after few minutes in contact with SUV suspension (Figure 1B). Similar round structures have already been seen by AFM for other lipid mixtures.16,32 The bright line in Figure 1B indicates where the height cross section (1G) was taken. The step height of 5 nm is in good agreement with DMPC/DPPC bilayers in extended conformation. The patch size increases with time leading to quasi-complete coverage of the mica surface with 7

some residual defects remaining clearly visible (Figure 1F). The height of the bilayer patches remains constant during SPB formation. During increasing coverage, lateral contacts between the patches induce the formation of nervure-like structures that are still visible after the final coverage is reached (Figure 1F). The nervure-like structures correspond to gaps which appear to be a few angstroms less deep than the bilayer height, but the measurement was not deconvoluted with respect to tip dimensions. The SPB formation was also studied by in situ AFM for a mixture of DMPC/DPPC in a ratio 70/30 (Figure 2). In this case, it is difficult to obtain good images during the phase of individual lipid bilayer patches. After reaching a critical surface coverage (Figure 2B and 2C), imaging becomes easier again (Figure 2D, 2E, and 2F). It should be noted that the nervure-like structures do not change significantly between 30 and 50 min (images not shown). The results on a considerably faster formation of a “complete” lipid bilayer on mica are in agreement with the reduced Tm of the 70/30 mixture (29 °C) in comparison with the 50/50 mixture (33 °C). It shows, however, that vesicle rupture and bilayer fusion are possible on mica at temperature slightly below Tm. For the present work, we have chosen mixtures for which the ambient temperature at which AFM experiments were carried out falls between the pretransition temperature and the main transition temperature of the lipid mixture in the bulk phase. In view of the fact that the transition temperatures for SUVs and for supported bilayers are slightly reduced in comparison to the bulk transition temperature, it was to be expected that the lipid mobility of the mixtures investigated was not too small to allow the formation of a supported bilayer from a vesicle phase even below the main transition. Consequently, the rate of bilayer formation of the 70/30 mixture is faster than the rate for the 50/50 mixture. A complete coverage is observed at t ) 34 min for the ratio 70/30 (Figure 2F), while in case of the 50/50 ratio a second injection of vesicles is needed. As a consequence of this observation, mixtures with a higher DPPC content were not tested. The height profile in Figure 2G shows a membrane thickness of 5 nm, which is similar to the one observed for mixture 50/50. The height resolution of the AFM images does not allow to determine the difference of the average heights of the bilayers of the different lipid mixtures, which would be expected to be slightly more than a few Å. It also prevents a more detailed interpretation with respect to an eventual enrichment of one of the lipids in the SPB in comparison to the original mixture or to an eventual phase separation on the surface. There are fewer defects in the final lipid bilayer for the 70/30 mixture than for the 50/50 mixture. Lateral contact between the lipid patches also leads to nervure-like structures that are somewhat larger in the 70/ 30 case (Figure 2F). Since AFM is a local probe and produces images of selected areas only, it is helpful to investigate the same systems by a method that integrates over macroscopic areas and that has a superior temperature control and resolution. A convenient alternative method for obtaining such data is the dissipation enhanced quartz crystal microbalance (QCM8

D (Q-Sense, Go¨teborg, Sweden)), which has been used before to monitor SPB formation. It has been described in detail by Rodahl et al.34 in 1995 and only a short outline is presented here. In addition to the well-known changes in resonance frequency, ∆f, with increasing adsorbed mass, it is possible to estimate viscoelastic data of the adsorbed layer by measuring the energy dissipation, ∆D. The fluid cell is temperature controlled to a precision of 0.05 °C and has a baseline stability over several hours. AT-cut quartz crystals (14 mm in diameter, Q-Sense) with a fundamental frequency of 5 MHz were used in the present study. In addition to the fundamental frequency, the overtones at the third, fifth, and seventh harmonic corresponding to 15, 25, and 35 MHz, respectively, are sampled as well. Crystals used for the present studies were coated with a ∼100 nm thick SiO2 film and used as received after cleaning. Cleaning of the crystals is achieved by successive rinses in water and ethanol followed by an UV/ozone treatment. Experiments start with two injections of 0.5 mL of buffer into the measurement cell (internal volume of 50 µL). At t ) 0, 0.5 mL of the SUV suspension is injected and changes in ∆f and ∆D are continuously recorded. A final rinse with buffer after stabilization of the signal (20-60 min) is used to test for weakly adsorbed material. The final rinse revealed that in all cases reported here no material was removed and therefore one can speak of stable adsorbed species. At first, the influence of the temperature on the formation of SPBs was studied by QCM-D for a pure lipid, DMPC, having a melting temperature of 24 °C (Figure 3A and A*). At 33 °C, which is 9 °C above the Tm, QCM-D signals shows the expected ∆f and ∆D curves for a SPB formation (Figure 3A and 3A*). Until t ) 3 min, the adsorption of vesicles dominates the observed variables (falling ∆f/n and rising ∆D). The regime from t ) 3 min to t ) 9 min is dominated by vesicle rupture and bilayer formation, as seen by rising ∆f and falling in ∆D. The final values of ∆f ) 25 Hz and ∆D ) 0.3 × 10-6, which do not change upon rinsing in buffer, indicate the formation of a complete and rather rigid lipid bilayer. At 24 °C, which corresponds to the phase transition temperature in bulk, a similar behavior is observed in frequency and in dissipation, also indicating the formation of a complete supported bilayer. However, the required time for obtaining a complete coverage of the surface is somewhat longer. The complete process takes 9 min at 33 °C and more than 30 min at 24 °C. Figures 3A and 3A* also show the small experimental error between the different harmonics. To keep a maximum of clarity, figures 3B, 3B*, 3C, and 3C* show the data of the third harmonic at 15 MHz only. In the following, we change the transition temperature of the bilayer by changing its composition. While pure DMPC has its transition temperature at 24 °C, it is the most fluid system at ambient conditions. Adding an increasing amount of DPPC, which transition temperature is 41 °C, reduces the fluidity as the transition temperature increases. As a consequence, it is to be expected that the kinetics of vesicle fusion and SPB formation is slow. While the experiments carried out at 37 °C are above the transition temperature of DMPC and all mixtures, the behavior of all three systems is Nano Lett., Vol. 4, No. 1, 2004

Figure 3. Influence of temperature on SPB formation on SiO2 displayed as ∆f(t) (left) and ∆D(t) (right) for pure DMPC (A and A*), DMPC/DPPC 70/30 (B and B*), and DMPC/DPPC 50/50 (C and C*). While all three harmonics are depicted for DMPC, only the third overtone is represented for DMPC/DPPC mixtures (70/30 and 50/50).

indistinguishable. In contrast, experiments carried out at 24 °C are below or close to the phase transition temperature, and thus the process of vesicle rupture and bilayer formation are slowed. For the mixture 50/50, the curves of ∆f/n and ∆D at 24 °C indicate that vesicles adsorb fast but rupture and fusion is slowed and that the transformation of vesicles to SPBs stays incomplete. A high value of ∆f/n and an increasing ∆D indicate that the adsorbed layer contains more mass and dissipates more energy than a closed bilayer. In contrast to the AFM experiments, we did not attempt to inject SUVs a second time to complete bilayer formation. We conclude from the QCM-D results that the temperature has a fundamental role in SPB formation especially for promoting vesicle rupture. Kasemo et al. have shown recently a strong effect of temperature on SPB formation kinetics above the phase transition temperature of PC-vesicles and Nano Lett., Vol. 4, No. 1, 2004

in the absence of calcium.20 By increasing the temperature, a lower coverage of intact vesicles is required to induce vesicle rupture. However, in the present work, the temperature effect observed for DMPC (Figure 3A) is less pronounced, which seems to be caused by the presence of calcium ions that modify the mechanism of vesicle rupture.31 It is often assumed that it is necessary to work above the melting temperature to form SPBs by vesicle fusion. However, as described above, we observe SPB formation at 24 °C, below Tm, for both DMPC/DPPC mixtures by AFM. Our observation that bilayer formation was observed to go to completion at 24 °C by AFM but that it was severely hampered by QCM-D at 24 °C merits further explanation. Beckmann et al. have already shown low temperature formation for a single lipid (C15PC).29 In fact, the two complementary techniques differ by the use of different 9

surfaces for AFM and QCM-D. Indeed, mica and silicon oxide surfaces exhibit different roughness, composition, crystallinity, and hydrophilic properties.20,35,36 From literature, experimental results suggest a different SPB formation mechanism on these two different surfaces. On mica, vesicle rupture occurs spontaneously, especially in the presence of divalent ions,32 while on SiO2, a critical coverage is observed by AFM and QCM-D before the rupture is induced.31 This way, this rupture process seems to be slower on the silicon oxide surface, allowing kinetic study of SPB formation. We assume that a potential heating effect of the laser passing through the solution in the AFM cell cannot be excluded, but it is estimated to be negligible. In this paper, we have demonstrated that using complementary surface analytical tools such as AFM and QCM-D is necessary to understand in situ kinetics of SPB formation close to the phase transition temperature of lipid mixtures where the observation of intermediate structures becomes possible. Sequential AFM images, done below the Tm, display transitional nanoscopic objects in an early stage that are growing in time to a quasi-complete coverage of the surface. However, QCM-D measurements performed in the same experimental conditions show the coexistence of SPBs and intact vesicles on the surface, while above the Tm SPBs are formed. A comparison with results obtained on pure DMPC demonstrates the importance of the temperature during SPB formation with respect to the Tm of the lipid mixtures. Vesicle rupture is promoted at temperatures higher than Tm, while this process is slowed at temperatures lower than Tm. DMPC/ DPPC mixtures illustrate perfectly this phenomenon. A correlation between AFM and QCM-D results has been possible for a kinetic aspect related to the phase transition temperature by taking into account the nature of the surfaces used. Acknowledgment. We acknowledge financial support from the University Louis Pasteur, the CNRS, and La Re´gion Alsace (contract 01/810). We thank Prof. Alain Brisson and Dr. Ralf Richter (IECB, University of Bordeaux 1) for helpful discussions. References (1) Decher, G.; Hong, J.-D. Makromol. Chem., Macromol. Symp. 1991, 46, 321-327. (2) Decher, G. Science 1997, 277, 1232-1237.

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(3) Decher, G.; Schlenoff, J. B. Multilayer Thin Films: Sequential Assembly of Nanocomposite Materials; Wiley-VCH: Weinheim, 2002. (4) Lvov, Y.; Essler, F.; Decher, G. J. Phys. Chem. 1993, 97, 1377313777. (5) Sukhorukov, G. B.; Donath, E.; Moya, S.; Susha, A. S.; Voigt, A.; Hartmann, J.; Mohwald, H. J. Microencapsul. 2000, 17, 117-185. (6) Zhang, L.; Longo, M. L.; Stroeve, P. Langmuir 2000, 16, 50935099. (7) Singer, S. J.; Nicolson, G. L. Science 1972, 175, 720-731. (8) Almeida, P. P. F.; Vaz, W. L. C. Structure and Dynamic of Membranes; Elsevier: Amsterdam, 1995. (9) Sackmann, E. Science 1996, 271, 43-48. (10) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105-103. (11) Bayerl, T. M.; Bloom, M. Biophys. J. 1990, 58, 357-362. (12) Salamon, Z.; Huang, D.; Cramer, W. A. Biophys. J. 1998, 75, 17841885. (13) Keller, C. A.; Kasemo, B. Biophys. J. 1998, 75, 1397-1402. (14) Steinem, C.; Janshoff, A.; Ulrich, W.-P.; Sieber, M.; Galla, H.-J. Biochim. Biophys. Acta, Biomembr. 1996, 1279, 169-180. (15) Cheng, Y.; Boden, N.; Bushby, R. J.; Clarkson, S.; Evans, S. D.; Knowles, P. F.; Marsh, A.; Miles, R. E. Langmuir 1998, 14, 839844. (16) Jass, J.; Tja¨rnhage, T.; Puu, G. Biophys. J. 2000, 79, 3153-3163. (17) Reviakine, I.; Brisson, A. Langmuir 2000, 16, 1806-1815. (18) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 25542559. (19) Salafsky, J.; Groves, J. T.; Boxer, S. G. Biochemistry 1996, 35, 14773-14781. (20) Reimhult, E.; Ho¨o¨k, F.; Kasemo, B. Langmuir 2003, 19, 1681-1691. (21) Lingler, S.; Rubinstein, I.; Knoll, W.; Offenha¨user, A. Langmuir 1997, 13, 7085-7091. (22) Cassier, T.; Sinner, A.; Offenha¨user, A.; Mo¨hwald, H. Colloids Surf. B: Biointerfaces 1999, 15, 215-225. (23) Bentz, J.; Duzgune, N.; S., N. Biochemistry 1983, 22, 3320-3330. (24) Garidel, P.; Blume, A. Biochim. Biophys. Acta 1998, 1371, 83-95. (25) Garidel, P.; Blume, A. Eur. Biophys. J. 2000, 28, 629-638. (26) Heimburg, T. Biophys. J. 2000, 78, 1154-1165. (27) Garidel, P.; Johann, C.; Blume, A. Biophys. J. 1997, 72, 21962210. (28) Linseisen, F. M.; Hetzer, M.; Brumm, T.; Bayerl, T. M. Biophys. J. 1997, 72, 1659-1667. (29) Beckmann, M.; Nollert, P.; Kolb, H.-A. J. Membr. Biol. 1998, 161, 227-233. (30) Reimhult, E.; Ho¨o¨k, F.; Kasemo, B. J. Chem. Phys. 2002, 117, 74017404. (31) Richter, R.; Mukhopadhyay, A.; Brisson, A. Biophys. J. 2003, 85, 3035-3047. (32) Egawa, H.; Furusawa, K. Langmuir 1999, 15, 1660-1666. (33) Seantier, B.; Breffa, C.; Fe´lix, O.; Decher, G. in preparation. (34) Rodahl, M.; Ho¨o¨k, F.; Krozer, A.; Brzezinki, P.; Kasemo, B. ReV. Sci. Instrum. 1995, 66, 3924-3930. (35) Richter, R.; Brisson, A. Langmuir 2003, 19, 1632-1640. (36) Iler, R. K. The Chemistry of Silica: Solubility, Polymerization, Colloid and Surface Properties and Biochemistry; John Wiley & Sons: New York, 1979.

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Nano Lett., Vol. 4, No. 1, 2004