Formation of Substrate-Supported Membranes from Mixtures of Long

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Formation of Substrate-Supported Membranes from Mixtures of Long- and Short-Chain Phospholipids Kenichi Morigaki,*,†,‡,§ Shigeki Kimura,§ Keisuke Okada,‡,§ Takashi Kawasaki,§ and Kazunori Kawasaki§ †

Research Center for Environmental Genomics and ‡Graduate School of Agricultural Science, Kobe University, Rokkodaicho 1-1, Nada, Kobe 657-8501 Japan § National Institute of Advanced Industrial Science and Technology (AIST), Ikeda 563-8577, Japan S Supporting Information *

ABSTRACT: We studied the formation of substrate-supported planar phospholipid bilayers (SPBs) on glass and silica from mixtures of long- and short-chain phospholipids to assess the effects of detergent additives on SPB formation. 1,2-Hexyanoylsn-glycero-3-phosphocholine (DHPC-C6) and 1,2-heptanoylsn-glycero-3-phosphocholine (DHPC-C7) were chosen as short-chain phospholipids. 1-Palmitoyl-2-oleol-sn-glycero-3phosphocholine (POPC) was used as a model long-chain phospholipid. Kinetic studies by quartz crystal microbalance with dissipation monitoring (QCM-D) showed that the presence of short-chain phospholipids significantly accelerated the formation of SPBs. Rapid rinsing with a buffer solution did not change the adsorbed mass on the surface if POPC/DHPC-C6 mixtures were used below the critical micelle concentration (cmc) of DHPCC6, indicating that an SPB composed of POPC molecules remained on the surface. Fluorescence microscopy observation showed homogeneous SPBs, and the fluorescence recovery after photobleaching (FRAP) measurements gave a diffusion coefficient comparable to that for SPBs formed from POPC vesicles. However, mixtures of POPC/DHPC-C7 resulted in a smaller mass of lipid adsorption on the substrate. FRAP measurements also yielded significantly smaller diffusion coefficients, suggesting the presence of defects. The different behaviors for DHPC-C6 and DHPC-C7 point to the dual roles of detergents to enhance the formation of SPBs and to destabilize them, depending on their structures and aggregation properties.

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INTRODUCTION Substrate-supported planar phospholipid bilayers (abbreviated as SPBs) are being studied extensively as a model cellular membrane.1,2 The most commonly used process for generating SPBs is the spontaneous adsorption and rupture of phospholipid vesicles on the substrate surface (vesicle spreading).3,4 Although it is a convenient self-assembly process, its applicability is limited to a relatively small range of phospholipids and substrate materials.5 In particular, it is generally difficult to incorporate membrane proteins into SPBs via vesicle spreading.6 An alternative method of vesicle spreading is the use of detergents, which can solubilize both lipids and proteins from biological membranes. There have been a number of studies that utilized lipid/detergent mixtures to attach lipid layers to a solid surface.7−12 Although the solubilization and reconstitution of membrane proteins using detergents have a long history of studies and a large accumulation of knowledge and protocols,13,14 the selfassembly of lipid/detergent mixtures at the solid/liquid interface is not fully understood and its application to the generation of SPBs is still limited.15 It would be advantageous if we could apply lipid/detergent self-assembly as a complementary approach to vesicle spreading. To obtain insight into the formation of SPBs from lipid/detergent mixtures, we studied the adsorption of membranes onto a solid surface from mixtures of long- and short-chain phospholipids. We chose 1,2© 2012 American Chemical Society

hexyanoyl-sn-glycero-3-phosphocholine (DHPC-C6) and 1,2heptanoyl-sn-glycero-3-phosphocholine (DHPC-C7) as model detergents in part because short-chain phospholipids are regarded as a mild detergent that can solubilize and reconstitute membrane proteins, retaining their native structures and functions (Figure 1).16,17 We were also inspired by the disklike structure formed from mixtures of long- and short-chain phospholipids (bicelles) that could be aligned in a magnetic field.18−21 We conducted kinetic and microscopic studies of membrane adsorption by using quartz crystal microbalance with dissipation monitoring (QCM-D) and total internal reflection fluorescence microscopy (TIR-FM). In the microscopic studies, we used micropatterned polymeric phospholipid bilayers composed of 1,2-bis(10,12-tricosadiynoyl)-sn-glycero3-phosphocholine (DiynePC) as a preformed matrix for incorporating POPC/DHPC mixtures. (Note that we use the term DHPC when referring to both DHPC-C6 and DHPCC7.) Polymerized bilayers are resistant to solubilization in the presence of detergents and provide matrices with a well-defined thickness (ca. 4.5 nm).22−24 We evaluated the lateral mobility of lipid molecules as a basic physicochemical property of formed SPBs. We compared the SPB formation in the presence Received: February 17, 2012 Revised: May 13, 2012 Published: May 16, 2012 9649

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solvent (30−45 min), the monolayer was compressed to a surface pressure of 34 mN/m. While the surface pressure remained constant, the monolayer was transferred to clean substrates. The first monolayer was deposited by dipping and withdrawing the substrate vertically (LB method). The second leaflet was deposited onto the hydrophobic surface of the first monolayer by pressing the substrate horizontally through the monolayer at the air/water interface and dropping it into the subphase (LS method). After the deposition of the second monolayer, the substrates were collected from the trough and stored in deionized water (in the dark) for polymerization. DiynePC bilayers were polymerized by UV irradiation using a mercury lamp (UVE502SD, Ushio, Tokyo, Japan) as the light source. The applied UV intensity and dose were typically 10 mW/cm2 and 5 J/cm2 at 254 nm, respectively. Desired micropatterns were imposed in DiynePC bilayers by illuminating the sample through a mask (a quartz slide with a patterned chromium layer coating) that was placed directly on the monomeric DiynePC bilayer. After UV irradiation, nonpolymerized DiynePC molecules were removed from the substrate surface by immersion in a 0.1 M sodium dodecylsulfate (SDS) solution at 30 °C for 30 min and rinsing with deionized water extensively. The polymerized DiynePC substrates were stored in deionized water in the dark at 4 °C for the experiments. QCM-D Measurements. QCM-D measurements were performed by using a Q-Sense D300 system with a QAFC 302 axial flow chamber (Q-Sense, Göteborg, Sweden). Quartz crystals with a thin SiO2 layer were used as the sensors (Q-sense, Göteborg, Sweden). For the measurements, the sensor crystal was oscillated at its resonance frequency of 5 MHz and at three harmonics (15, 25, and 35 MHz), and was monitored for the shifts in frequency (Δf) and dissipation (ΔD). The interval for data acquisition was 0.4 s. The mounted QCM sensor crystal was first equilibrated with a degassed buffer solution at 21.8 °C. The buffer solution was subsequently replaced with POPC/ DHPC suspensions. Fluorescence Microscopy Observation. Fluorescence microscopy observations of POPC/DHPC aggregates were performed by using an Olympus IX81 inverted microscope with a 60x PlanApo TIRFM oil-immersion objective (NA 1.45, Olympus, Tokyo, Japan). An argon ion laser (excited at 488 nm) was used as the light source for TIR-FM, and a xenon lamp (AH2-RX-T, Olympus) was used for epifluorescence. Polymeric DiynePC was observed by TIR-FM using a filter set for green fluorescence (termed TIR-FM-G herein), and TRPE was observed either by TIR-FM using a filter set for red fluorescence (termed TIR-FM-R herein) or the Olympus U-MWIY2 filter set (excitation wavelength 545−580 nm, emission wavelength >610 nm, abbreviated as WIY) for epifluorescence. Fluorescence images were obtained with a CCD camera (C4742-95-12ERG, Hamamatsu Photonics, Hamamatsu, Japan). For the observation of SR membranes, we used an Olympus BX51WI upright microscope with a 20× water-immersion objective (NA 0.95, Olympus). A filter set having an excitation wavelength range of 470−490 nm and an emission wavelength range of 510−550 nm (abbreviated as NIBA) was used. Fluorescent images were collected with a CCD camera (DP30BW, Olympus). The fluidity of SPBs was determined by the fluorescence recovery after photobleaching (FRAP) analysis using the boundary profile evolution (BPE) method.26 A rectangular area was photobleached by illumination through a slit, and the fluorescence profiles at the boundary region between the bleached and nonbleached areas were observed with time. (The photobleached area was sufficiently large so that the boundary could be regarded as 1D.) Examples of boundary profiles are given in the Supporting Information (Figure S4). The collected boundary profiles were fitted to a Gaussian error function by using the Origin program (OriginLab Corporation, MA) to determine the diffusion depth w that is defined as

Figure 1. Chemical structures of POPC, DHPC-C6, and DHPC-C7.

of DHPC-C6 and DHPC-C7 to highlight the dual roles of detergents in the reconstitution process.

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MATERIALS AND METHODS

Materials. 1,2-Hexyanoyl-sn-glycero-3-phosphocholine (DHPCC6), 1,2-heptanoyl-sn-glycero-3-phosphocholine (DHPC-C7), 1-palmitoyl-2-oleol-sn-glycero-3-phosphocholine (POPC), and 1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine (DiynePC) were purchased from Avanti Polar Lipids (Alabaster, AL). Texas Red 1,2dihexadecanoyl-sn-glycero-phosphoethanolamine (TR-PE) and 3,3′dioctadecyloxacarbocyanine perchlorate (DiO) were purchased from Molecular Probes (Eugene, OR). Deionized water used in the experiments was ultrapure Milli-Q water (Millipore) with a resistance of 18.2 MΩ cm. It was used to clean substrates and prepare buffer solutions (0.02 M Hepes buffer with 0.15 M NaCl, pH 7.4) and for all other experiments. Substrate Cleaning. Microscopy glass slides and coverslips (Matsunami, Osaka, Japan) were used as substrates for fluorescence microscopy observation. The substrates were cleaned with a commercial detergent solution, 0.5% Hellmanex/water (Hellma, Mühlheim, Germany), for 20 min under sonication, rinsed with deionized water, treated in a solution of NH4OH (28%)/H2O2 (30%)/H2O (0.05:1:5) for 10 min at 65 °C, rinsed extensively with deionized water, and then dried in a vacuum oven for 30 min at 80 °C. Before use, these substrates were further cleaned via UV/ozone treatment for 20 min (PL16-110, Sen Lights Corporation, Toyonaka, Japan). Preparation of POPC/DHPC Mixed Aggregates. POPC dissolved in chloroform was dried in a stream of nitrogen and subsequently evaporated at least for 4 h in a vacuum desiccator. The dried lipid film was hydrated in a buffer solution containing DHPC of the required concentration. (The POPC concentration was 1 mM for all measurements.) The resulting suspensions of POPC/DHPC mixed aggregates were put through five freeze/thaw cycles. The solution was stored in the dark at 4 °C overnight and vortex mixed before use. Preparation of Patterned DiynePC Bilayers. A detailed description of the patterned bilayer formation using polymerized DiynePC is given in previous papers.24,25 Briefly, bilayers of monomeric DiynePC were deposited onto substrates from the air/ water interface by the Langmuir−Blodgett (LB) and subsequent Langmuir−Schaefer (LS) methods using a Langmuir trough (HBMAP, Kyowa Interface Science, Asaka, Japan). The temperature of the subphase (deionized water) was controlled to 16 °C by circulating thermostatted water. Monomeric DiynePC was spread onto the subphase from a chloroform solution. After the evaporation of the

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F(x , t ) − Fbleached ⎛ x − xb ⎞ ⎟ + 1 = erf⎜ ⎝ 2w ⎠ Funbleached − Fbleached

w= 9650

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where D is the diffusion coefficient and t is the time after photobleaching. F(x, t) represents the fluorescence intensity at a given time t and position x in the vertical axis with respect to the boundary. xb is the boundary position between the bleached and unbleached areas. Fbleached and Funbleached are fluorescence intensities in the bleached and unbleached areas that are far enough from the boundary, respectively. From the obtained w values, w2 values were plotted versus time t. The diffusion coefficient D of fluorescent molecules was determined from the slope of the linear dependency. (An advantage of the FRAP-BPE method over the more conventional FRAP technique based on pinhole photobleaching is the fact that the results are not affected by the timing of the initial measurement.) Electron Microscopy Observation. An aliquot of sample was put on a copper grid (400 mesh) coated with collodion and negatively stained by rinsing with 1% (w/v) uranyl acetate. The sample grid was dried in air, followed by the deposition of carbon vapor under vacuum. The morphology of POPC/DHPC mixed aggregates was examined with a transmission electron microscope (Tecnai G2 F20, FEI, Hillsboro, OR) operated at 120 kV.

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RESULTS AND DISCUSSION Monitoring Bilayer Adsorption by QCM-D. The adsorption process of mixed POPC/DHPC aggregates on a silica surface was monitored by QCM-D. QCM-D monitors the mass of adsorbed molecules on the surface as changes in the resonance frequency (Δf).27 It can also assess the viscoelasticity of the adsorbed material by measuring the damping of oscillation (dissipation ΔD). Figure 2 shows the results for the varied ratio of POPC and DHPC-C6 (q = [POPC]/ [DHPC-C6]). We held the concentration of POPC constant at 1 mM and varied the concentration of DHPC-C6 (except for q = 0.025, in which [POPC] = 0.5 mM and [DHPC-C6] = 20 mM, and q = 0). The formation of a planar bilayer from pure POPC vesicles (q = ∞) proceeded in two phases, as reported in the literature.27 In the first phase, the adsorption of vesicles on the surface occurred, resulting in the decrease of Δf and the increase of ΔD. In the second phase, vesicles ruptured and reorganized into a planar bilayer on the substrate surface. (It should be noted that we did not change the sizes of aggregates by processes such as extrusion or sonication. It is rather surprising that SPB formation was observed from nonextruded POPC vesicles, but presumably the vesicular suspension contained a heterogeneous mixture of large multilamellar vesicles and smaller unilamellar vesicles, enabling the transformation into an SPB.) The presence of DHPC-C6 dramatically accelerated the planar bilayer formation, as indicated by the shortened time for reaching the final values of Δf and ΔD (Figure 2A). The degree of acceleration was enhanced as the q values became smaller. Also for large q values (q = 100, 10 (i.e., 1, 10% of DHPC with respect to POPC)), the time required to reach the Δf minimum and ΔD maximum decreased, suggesting that the rupture of adsorbed vesicles occurred more rapidly. As the q values decreased further (more DHPC-C6 with respect to POPC), the accelerated formation of supported membranes became more prominent. The overshooting of Δf and ΔD was not observed for q values smaller than 0.1. The lack of overshooting indicates the rapid transformation of adsorbed POPC/DHPC-C6 aggregates into a planar bilayer.28 The observed acceleration of planar membrane formation in the presence of DHPC-C6 should stem from the following factors. The first factor is the change in membrane rigidity (bending elasticity). It has been reported that amphiphilic molecules that increase the spontaneous curvature of the membrane (e.g., lysolecithin) reduces the kinetic barrier of vesicle rupture and reorganization on the

Figure 2. (A) QCM-D profiles of the membrane adsorption process on the SiO2 surface for POPC/DHPC-C6 mixtures with varied compositions (q = [POPC]/ [DHPC-C6]). The concentration of POPC was kept constant at 1 mM, and the concentration of DHPCC6 was changed, except for q = 0.025, in which [POPC] = 0.5 mM and [DHPC-C6] = 20 mM. The frames on the right side are expansions of those on the left, showing the adsorption behaviors in the initial stage. (B) Final ΔD and Δf values of the QCM-D measurements on SiO2 (before rinsing) were plotted vs the POPC/ DHPC-C6 compositions.

substrate surface.29 At a high concentration of DHPC-C6, the transformation of the membrane structure should also play an important role. Electron microscopy observation of POPC/ DHPC-C6 mixtures have revealed that spherical vesicles transformed into corrugated disklike membranes as the concentration of DHPC-C6 increased (Figure 3). It should be noted that the sample preparation process for the negative staining electron microscopy could affect the aggregation structures. However, the observed morphologies do differ among the three samples with different q values. In fact, we observed normal vesicular structures for q = ∞, as expected, whereas significant shape changes occurred for samples with smaller q values. This observation coincides with the tendencies reported in the literature and also gives insights into the mechanism by which aggregation structures affect the selfassembly at the interface.21 It is therefore quite sensible to assume that these structural changes should alter the kinetic behaviors of membrane transformation. The second factor is changes in the mean sizes of aggregates. Smaller aggregates can diffuse more rapidly onto the surface, accelerating the membrane formation. The addition of DHPC-C6 to the POPC vesicle suspension is expected to solubilize POPC 9651

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Figure 3. Representative electron microscope images of POPC/ DHPC-C6 mixed aggregates. The compositions of the samples were (A) q = ∞, (B) q = 10, and (C) q = 5, respectively. The bars indicate 50 nm.

Figure 4. QCM-D profiles during the rinsing process of POPC/ DHPC-C6 mixtures having varied compositions. An aqueous solution containing POPC (1 mM) and DHPC-C6 (varied concentrations) was rapidly replaced with a buffer solution. The time point of the solution exchange is indicated by an arrow.

bilayers, forming smaller structures as evidenced by the electron microscopy observation (Figure 3) and the reduction of turbidity (Supporting Information, Figure S1). Also for intermediate mixtures we observed a substantial reduction of turbidity, presumably because of the reduction of the mean aggregation sizes. van Dam et al. have reported that mixtures of long- and short-chain phospholipids formed smaller micellar structures for a lower q value and that the size increased drastically as the total lipid concentration decreased.21 Having a higher concentration of DHPC-C6 decreases q and increases the total lipid concentration at the same time, both of which should have the effect of reducing the aggregation sizes. The final values of ΔD and Δf are plotted as a function of the q values in Figure 2B. These values represent the mass and dissipation of adsorbed lipid layers that are in equilibrium with lipid/detergent mixtures in the solution. For the q values above 0.1, Δf was 25−26 Hz and ΔD was relatively small (1 μm). Forth, polymerized DiynePC emits fluorescence that is due to its conjugated polymer backbone, making the observation of the surface possible in the absence of other fluorescent probes. These features render patterned polymeric DiynePC bilayers a useful matrix for the incorporation and observation of lipid/ detergent aggregates on the surface. Figure 6 shows the fluorescence micrographs for the incorporation of POPC/ DHPC-C6 (q = 0.1: [POPC] = 1 mM, [DHPC] = 10 mM, [TR-PE] = 0.01 mM) into a polymeric DiynePC bilayer matrix. Figure 6A is a TIR-FM observation of a polymeric DiynePC bilayer in the shape of a grid (the size of the square corrals was 20 μm). Figure 6B is a TIR-FM observation of POPC/DHPCC6 membranes in the presence of aggregates in the bulk solution (same position as in Figure 6A). We observed the homogeneous fluorescence of TR-PE in the corrals. Figure 6C shows the TIR-FM observation of incorporated bilayers after rinsing POPC/DHPC-C6 in the bulk solution with a buffer solution. The observed features of the membrane were unchanged from those in Figure 6B. Figure 6D is an epifluorescence observation at the same location as in Figure 6C, which confirms that there are no residual aggregates in the solution. By locally photobleaching the fluorescence in the membrane and observing its recovery with time, we could

confirm that the membranes incorporated into the corrals were continuous and fluid (vide infra). These observations support the results from the QCM-D experiments that a planar bilayer membrane was formed on the surface even at a high concentration of DHPC-C6. The formation of homogeneous SPB was microscopically observed also from POPC/DHPC-C7 if the concentration of DHPC-C7 was below the cmc, although many aggregates were also observed on a polymerized bilayer (Supporting Information, Figure S3). For a quantitative evaluation of the lateral mobility of lipid molecules, we applied the FRAP-BPE method.26 The obtained diffusion coefficients were 1.50 ± 0.27, 1.44 ± 0.16, and 0.96 ± 0.13 μm2/s for SPBs formed from POPC, POPC/DHPC-C6 ([DHPC-C6] = 10 mM), and POPC/DHPC-C7 ([DHPC-C7] = 1 mM), respectively. The values obtained for POPC were in the range of the diffusion coefficients reported in the literature, which is rather diverse because of different measurement conditions.26,35 The diffusion coefficient of POPC/DHPC-C6 was also comparable to that of POPC. However, the diffusion coefficient of POPC/DHPC-C7 was significantly lower. The lowered mobility of molecules also suggests the presence of defects in the SPB. Roles of Detergents in SPB Formation. The different results of SPB formation in the presence of DHPC-C6 and DHPC-C7 point to the dual roles of detergents in enhancing the formation of SPBs and in destabilizing the bilayer structures. The acceleration of SPB formation can be ascribed to the reduced kinetic barrier of vesicle rupture and reorganization on the substrate surface.29 At higher concentrations, the transformation of membranes into disklike structures should also play an important role. However, an obvious drawback of using detergents is the disruption of bilayer structures as a result of solubilization. Destabilization effects are generally greater as the detergent concentration is increased. Bilayer membranes are mostly disrupted if the detergent concentration is above its cmc. At the same time, working below the cmc does not necessarily lead to the 9653

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(Supporting Information, Figure S5), suggesting the possibility that we could also modulate the membrane adsorption for biological samples using a relatively mild detergent. The structurally controlled reconstitution of biological membranes on the solid substrate represents the next challenge to be addressed in parallel with the development of our knowledge of the lipid/detergent self-assembly process at the interface.

formation of intact bilayer structures. SPBs obtained in the presence of 10 mM DHPC-C6 were mostly comparable to those obtained from pure POPC vesicles. However, SPBs formed in the presence of 1 mM DHPC-C7 resulted in defects. Because the lipid/detergent ratio was generally greater for DHPC-C7 in the present study and a decreased mass was observed both below and above the cmc (Supporting Information Figure S2), the different observations between DHPC-C6 and DHPC-C7 should stem from their different affinities for bilayer membranes. Because DHPC-C6 has shorter alkyl chains than does DHPC-C7 by one methylene group, it has a lower affinity for bilayer membranes. Although DHPC-C6 is an integral part of the aggregation structure in the bulk solution, as suggested by the morphological changes observed by electron microscopy, it seems to have a less-significant influence on the bilayer structure on the solid substrate, as suggested by the QCM-D results. It should be noted that having a higher cmc does not always result in intact SPB formation as in the case of DHPC-C6. Octylglucoside, which has a higher cmc than DHPC-C6 (25.3 mM),36 did not result in intact SPBs if it was mixed with POPC (data not shown). Therefore, the formation of SPBs does not depend only on the hydrophilicity of detergent molecules; the structure of the molecules also plays an important role.

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ASSOCIATED CONTENT

S Supporting Information *

Turbidity of POPC/DHPC-C6 suspensions. QCM-D profiles of POPC/DHPC-C7 at different DHPC-C7 concentrations. Fluorescence micrographs of adsorbed bilayer membranes. Fluorescence recovery after photobleaching analysis using the boundary profile evolution (FRAP-BPE). Incorporation of biological membranes. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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CONCLUDING REMARKS We studied the formation of SPBs from mixtures of long- and short-chain phospholipids. The presence of DHPC-C6 accelerated the formation of SPBs, as monitored by QCM-D. The acceleration was attributed to the changes in the aggregation structure and the mechanical properties of the membranes. Rapid rinsing with a buffer solution did not deplete the membrane from the surface if the concentration of DHPCC6 was below its cmc. A fluorescence microscopy observation showed homogeneous SPB formation in the corrals between polymerized DiynePC. FRAP measurements also demonstrated the fluidity of the membranes, which is comparable to the fluidity of those formed from the vesicle spreading of POPC. Altogether, these results clearly support the formation of SPB in the presence of DHPC-C6. However, the addition of DHPCC7 to POPC vesicles resulted in SPBs with a lower adsorbed mass and smaller diffusion coefficients, suggesting the presence of defects. The self-assembly of the lipid−detergent mixture at the solid surface depends critically on factors such as the detergent structure, lipid concentration, and lipid/detergent ratio. There have been reports on the use of detergents for reconstituting model biological membranes on the surface. However, their applicability remains rather empirical, mainly because of the lack of a systematic comprehensive understanding of the self-assembly process at the interfaces. Our work has demonstrated that surface-specific analytical techniques such as QCM-D and TIR-FM are powerful tools in evaluating the reconstitution process with high sensitivity. In addition, micropatterned polymeric lipid bilayers also offer a useful scaffold for the analysis of surface-bound aggregation structures because they are robust, resistant toward nonspecific adsorption, and provide boundaries with a well-defined structure and thickness. The enhanced formation of SPBs in the presence of DHPC-C6 can potentially be extended to biological membranes. Our preliminary observation showed a significantly facilitated incorporation of sarcoplasmic reticulum (SR) membranes from rabbit skeletal muscle onto the solid substrate (glass slides) in the presence of DHPC-C6

ACKNOWLEDGMENTS We thank Ms. Saori Mori and Ms. Maki Koike for their assistance with the preparation of samples and the QCM-D measurements. We thank Mr. Tomoki Kato for his assistance with the electron microscopy observations. This work was supported by the Promotion Budget for Science and Technology (AIST Upbringing of Talent in Nanobiotechnology Course) from the Ministry of Education, Science, Culture and Sports (MEXT), the Grant-in-aid for scientific research from the Japan Society for the Promotion of Science (nos. 18510107 and 21023021), and the Sekisui Chemical Grant Program.

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REFERENCES

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dx.doi.org/10.1021/la300696z | Langmuir 2012, 28, 9649−9655