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Polymerized Lipid Bilayers on a Solid Substrate: Morphologies and Obstruction of Lateral Diffusion Takashi Okazaki,† Takehiko Inaba,† Yoshiro Tatsu,† Ryugo Tero,‡ Tsuneo Urisu,‡ and Kenichi Morigaki*,† Research Institute for Cell Engineering, National Institute of AdVanced Industrial Science and Technology (AIST), Ikeda 563-8577, Japan, and Institute for Molecular Science, Myodaiji, Okazaki 444-8585, Japan ReceiVed August 15, 2008. ReVised Manuscript ReceiVed October 22, 2008 Substrate supported planar lipid bilayers (SPBs) are versatile models of the biological membrane in biophysical studies and biomedical applications. We previously developed a methodology for generating SPBs composed of polymeric and fluid phospholipid bilayers by using a photopolymerizable diacetylene phospholipid (DiynePC). Polymeric bilayers could be generated with micropatterns by conventional photolithography, and the degree of polymerization could be controlled by modulating UV irradiation doses. After removing nonreacted monomers, fluid lipid membranes could be integrated with polymeric bilayers. Herein, we report on a quantitative study of the morphology of polymeric bilayer domains and their obstruction toward lateral diffusion of membrane-associated molecules. Atomic force microscopy (AFM) observations revealed that polymerized DiynePC bilayers were formed as nanometer-sized domains. The ratio of polymeric and fluid bilayers could be modulated quantitatively by changing the UV irradiation dose for photopolymerization. Lateral diffusion coefficients of lipid molecules in fluid bilayers were measured by fluorescence recovery after photobleaching (FRAP) and correlated with the amount of polymeric bilayer domains on the substrate. Controlled domain structures, lipid compositions, and lateral mobility in the model membranes should allow us to fabricate model membranes that mimic complex features of biological membranes with well-defined structures and physicochemical properties.
Introduction Substrate supported planar lipid bilayers (SPBs) are being studied as a model of biological membranes.1-4 One of the important features of SPBs is the possibility to generate micropatterned membranes on the substrate, which enables one to create designed arrays of biological materials for various applications, such as high throughput drug screening.5 A variety of approaches have been applied, including the use of prepatterned substrates,6-10 microcontact printing,11 microfluidics,12,13 and inkjet printers.5 We have previously developed a methodology for fabricating micropatterned SPBs by using polymerized phospholipid bilayers.14-16 The fabrication process comprises four steps, as illustrated in Figure 1: (A) formation of a monomeric bilayer on a solid substrate, (B) photolithographic polymerization by UV light, (C) removal of the nonreacted monomers, and (D) * Corresponding author. E-mail:
[email protected]. † National Institute of Advanced Industrial Science and Technology (AIST). ‡ Institute for Molecular Science.
(1) Sackmann, E. Science 1996, 271, 43–8. (2) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149–157. (3) Tanaka, M.; Sackmann, E. Nature 2005, 437, 656–663. (4) Scho¨nherr, H.; Degenhart, G. H.; Dordi, B.; Feng, C. L.; Rozkiewicz, D. I.; Shovsky, A.; Vancso, G. J. Organic and macromolecular films and assemblies as (bio)reactive platforms: From model studies on structure-reactivity relationships to submicrometer patterning. In Ordered Polymeric Nanostructures at Surfaces; Vancso, G. J., Reiter, G., Eds.; Springer: Berlin, 2006; Vol. 200, pp 169-208. (5) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124, 2394– 2395. (6) Groves, J. T.; Ulman, N.; Boxer, S. G. Science 1997, 275, 651–3. (7) Heyse, S.; Ernst, O. P.; Dienes, Z.; Hofmann, K. P.; Vogel, H. Biochemistry 1998, 37, 507–522. (8) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 2554–2559. (9) Jenkins, A. T. A.; Boden, N.; Bushby, R. J.; Evans, S. D.; Knowles, P. F.; Miles, R. E.; Ogier, S. D.; Scho¨nherr, H.; Vancso, G. J. J. Am. Chem. Soc. 1999, 121, 5274–5280. (10) Tanaka, M.; Wong, A. P.; Rehfeldt, F.; Tutus, M.; Kaufmann, S. J. Am. Chem. Soc. 2004, 126, 3257–3260. (11) Hovis, J. S.; Boxer, S. G. Langmuir 2000, 16, 894–897. (12) Kam, L.; Boxer, S. G. J. Am. Chem. Soc. 2000, 122, 12901–12902. (13) Ku¨nneke, S.; Janshoff, A. Angew. Chem., Int. Ed. 2002, 41, 314.
refilling the lipid-free regions with new lipid bilayers. As the polymerizable bilayer entities, we have used a diacetylene phospholipid, 1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine (DiynePC). The chemical structure and its polymerization scheme are given in Figure 1E. The polymerization proceeds as a 1,4-addition reaction, forming conjugated backbones of ene-yne bonds.17,18 The polymeric bilayer acts as a framework that supports embedded lipid membranes with defined boundaries. The embedded lipid membranes, on the other hand, retain some important characteristics of the biological membrane such as fluidity and are intended to be used as model systems of the biological membrane. The fact that the polymeric scaffolds and incorporated fluid membranes have the same bilayer structure gives some unique features to the model membrane. We have previously observed that incorporation of SPBs by vesicle fusion was significantly accelerated by the presence of preformed polymeric bilayers.19 Another important feature of the hybrid SPBs is the potential to regulate the mobility of membrane associated molecules using composite membranes of partially polymerized and fluid bilayers.16 For exploiting these potentials and utilizing the composite membranes in basic science and biomedical applications, it is important to evaluate their structural and physicochemical properties in depth. Therefore, we conducted a detailed study on the morphology of polymerized bilayer domains and their effects on the lateral mobility of lipid molecules within fluid bilayers. Atomic force microscopy (AFM) studies revealed that polymeric bilayers were composed of small domains of the order of 10 nm. (14) Morigaki, K.; Baumgart, T.; Offenha¨usser, A.; Knoll, W. Angew. Chem., Int. Ed. 2001, 40, 172–174. (15) Morigaki, K.; Baumgart, T.; Jonas, U.; Offenha¨usser, A.; Knoll, W. Langmuir 2002, 18, 4082–4089. (16) Morigaki, K.; Kiyosue, K.; Taguchi, T. Langmuir 2004, 20, 7729–7735. (17) Lopez, E.; O’Brien, D. F.; Whitesides, T. H. J. Am. Chem. Soc. 1982, 104, 305–307. (18) Bubeck, C.; Tieke, B.; Wegner, G. Ber. Bunsen-Ges. 1982, 86, 495–498. (19) Okazaki, T.; Morigaki, K.; Taguchi, T. Biophys. J. 2006, 91, 1757–1766.
10.1021/la802670t CCC: $40.75 2009 American Chemical Society Published on Web 12/09/2008
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Figure 1. (A-D) Schematic procedure of the construction of hybrid membranes composed of polymerized and fluid phospholipid bilayers. (E) Chemical structure of polymerizable diacetylene phospholipid, DiynePC, and its photopolymerization scheme: the marked acyl chains are linked upon polymerization, changing the orientation and expanding the DiynePC layer in the lateral direction.
The amount of polymeric bilayers on the surface could be modulated by changing the UV irradiation dose as measured by ellipsometry. After removing monomers and introducing fluid bilayers, the area fractions of polymeric and fluid bilayers were determined by fluorescence microscopy. Polymeric bilayer domains obstructed the lateral diffusion of lipids in fluid bilayers, depending on their density on the surface. We determined lateral diffusion coefficients of lipids and correlated them with the amount of polymeric bilayer domains by applying the percolation theory. The obtained quantitative information on the domain structures, lipid compositions, and lateral mobility defines the structural and physicochemical properties of the model membrane, and should facilitate their use in membrane biophysics studies as well as biomedical applications.
Materials and Methods Materials. Diacetylene phospholipid (1,2-bis(10,12-tricosadiynoyl)sn-glycero-3-phosphocholine (DiynePC)) and phosphatidylcholine from egg yolk (egg-PC) were purchased from Avanti Polar Lipids (Alabaster, AL). Texas Red 1,2-dihexadecanoyl-sn-glycero-phos-
Okazaki et al. phoethanolamine (TR-PE) was purchased from Molecular Probes (Eugene, OR). Sodium dodecyl sulfate (SDS) was purchased from Nacalai Tesque (Kyoto, Japan). Deionized water used in the experiments was ultrapure Milli-Q water (Millipore) with a resistance of 18.2 MΩ cm. It was used for cleaning substrates and preparing buffer solutions (0.01 M phosphate buffer with 0.15 M NaCl, pH 6.6) and as the subphase of the Langmuir monolayer. Substrate Cleaning. Microscopy glass slides, coverslips (Matsunami, Osaka, Japan), and boron doped (p-type) silicon (100) wafers covered with a native oxide layer (KN platz, Osaka, Japan) were used as substrates for fluorescence microscopy observation, AFM observation, and ellipsometric measurements, respectively. The substrates were cleaned first with a commercial detergent solution, 0.5% Hellmanex/water (Hellma, Mu¨hlheim, 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 by the UV/ozone treatment for 20 min (PL16-110, Sen Lights Corporation, Toyonaka, Japan). Deposition of Monomeric DiynePC Bilayers. 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 at 16 °C by using a circulating thermostatted water. Monomeric DiynePC was spread onto the subphase from a chloroform solution. After evaporation of the solvent (30-45 min), the monolayer was compressed to the surface pressure of 34 mN/m. While keeping the surface pressure constant, the monolayer was transferred onto cleaned 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 the polymerization. Polymerization of DiynePC Bilayers. Polymerization of DiynePC bilayers was conducted by UV irradiation using a mercury lamp (UVE-502SD, Ushio, Tokyo, Japan) as the light source. A closed system composed of a deionized water reservoir, a pump, and a cell (∼4 mL volume) was used. The water reservoir was depleted of oxygen by purging with argon. Deaerated water was circulated continuously with the pump through the polymerization cell. The cell had two walls on opposite sides, with one being the sample (the monomeric DiynePC bilayer was inside the cell) and the other being a quartz window through which UV light was illuminated. After sufficient circulation of deaerated water (typically for 15 min), the pump was stopped and the polymerization was started. The applied UV intensity was typically 10 mW/cm2 at 254 nm. The applied UV irradiation dose was varied by changing the exposure time. Desired micropatterns were imposed in DiynePC bilayers by illuminating the sample through a mask (a quartz slide with a patterned chromium layer coating) which was placed directly on the monomeric DiynePC bilayer. After UV irradiation, nonpolymerized DiynePC molecules were removed from the substrate surface by immersing in 0.1 M 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. Preparation of Lipid Vesicles. Vesicle suspensions of egg-PC containing 1 mol % TR-PE were prepared according to the following protocol. The lipids were mixed in a chloroform solution and then dried under a stream of nitrogen and subsequently evaporated at least for 4 h in a vacuum desiccator. The dried lipids were hydrated in a buffer solution overnight (the lipid concentration was 1 mM in the buffer solution). The resulting multilamellar vesicles were put through five freeze/thaw cycles. The vesicle suspensions were stored in the dark at 4 °C and extruded by using a LiposoFast extruder (Avestin, Ottawa, Canada) just before use, 10 times through a
Polymerized Lipid Bilayers on a Solid Substrate polycarbonate filter with 100 nm pores and subsequently 15 times through a polycarbonate filter with 50 nm pores. AFM Observation. AFM observations of polymeric bilayer surfaces were performed in deionized water by using the magneticAC mode with a PicoScan2500 instrument (Agilent Technologies, Inc. Santa Clara, CA). The collected images were treated with the scanning probe image processor (SPIP; Image Metrology A/S, Denmark) for plane correction and line profiling. The bilayer coverage was estimated with the ImageJ program (http://rsb.info.nih.gov/ij/). Fluorescence Microscopy Observation. For estimating the amount of incorporated fluid lipid membranes, microscopy observations were performed by using an OLYMPUS BX51WI upright microscope with a 60× water-immersion objective (NA 0.90, Olympus) and a xenon lamp (AH2-RX-T, Olympus). Polymeric DiynePC was observed by using the Olympus U-MNIBA2 filter set (excitation wavelength, 470-490 nm; emission wavelength, 510-550 nm; abbreviated as NIBA), and TR-PE was observed by using the Olympus U-MWIY2 filter set (excitation wavelength, 545-580 nm; emission wavelength, >610 nm; abbreviated as WIY). Fluorescence microscopic images were collected with a CCD camera (DP30BW, Olympus) and processed with the MetaMorph program (Molecular Devices, Sunnyvale, CA). Fluorescence Recovery after Photobleaching (FRAP) Analysis Using the Boundary Profile Evolution (BPE) Method. The fluidity of lipid bilayers within polymeric DiynePC matrices was determined by FRAP analysis using the BPE method, because it allows us to determine quantitative lateral diffusion coefficients without stateof-the-art experimental setups and it is applicable to a wide range of diffusion coefficients.20 Fluorescence images of TR-PE in a hybrid bilayer (no pattern was imposed in these experiments) were obtained by using the above-mentioned upright fluorescence microscope with a 60× water-immersion objective and the WIY filter set. Photobleaching through a rectangular slit was performed by keeping the shutter open for 5 or 10 s without a ND filter (full power of lamp). The photobleached area was sufficiently large for observing a boundary without being affected by the other three boundaries. After photobleaching, the changes in the fluorescence profiles at the boundary region between the bleached and unbleached areas were observed. The collected boundary profiles were fitted to a Gaussian error function by using the Origin program (OriginLab Corporation, Northampton, MA) for determining the diffusion depth w which is defined as
2
( )
F(x, t) - Fbleached x - xb ) erf +1 Funbleached - Fbleached 2w w ) √Dt
where D is the diffusion coefficient and t is the time after photobleaching. From obtained w values, the w2 values were plotted versus the time t. The diffusion coefficient D of fluorescent molecules was determined from the slope of linear dependency. Ellipsometric Measurement. Ellipsometric measurements were conducted in air by using an automated ellipsometer system DHAXA2Y (Mizojiri Opt. Co., Ltd.) with a He-Ne laser. Silicon wafers covered with a native oxide layer (KN platz, Osaka, Japan) were used as substrates. DiynePC bilayers were deposited by the LB/LS technique and homogeneously polymerized. After treating in an SDS solution, the residual polymer samples were dried under a stream of nitrogen just before the measurement. The refractive indices for silicon, silicon oxide, and DiynePC layers were assumed to be 3.875, 1.46, and 1.49, respectively. Quartz Crystal Microbalance with Dissipation (QCM-D) Measurement. QCM-D measurements were performed by using a Q-Sense D300 system with a QAFC 302 axial flow chamber (QSense, Go¨teborg, Sweden). Quartz crystals with a thin SiO2 layer were used as the sensors (Q-sense, Go¨teborg, Sweden). The sensor crystals were cleaned in 0.1 M SDS solution (immersed for 30 min at 30 °C), rinsed with deionized water, and dried under a nitrogen (20) Merzlyakov, M.; Li, E.; Hristova, K. Langmuir 2006, 22, 1247–1253.
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Figure 2. (A) AFM image (80 × 80 µm2) of a patterned polymeric bilayer after the SDS treatment. (B) Magnified image (1 × 1 µm2) of the boundary marked in (A). (C) Line profile measured along the line in (B).
stream. 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 the shifts of frequency (∆f) and dissipation (∆D) were monitored. 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 vesicle suspensions (lipid concentration 140 µM). Some sensors were coated with DiynePC bilayers. The bilayers were polymerized homogeneously with various UV irradiation doses (0.5, 1.0, 2.0, or 4.0 J/cm2), and nonreacted monomers were removed with 0.1 M SDS at 30 °C. The sensors with preformed polymeric DiynePC bilayers were stored in deionized water until they were used for the QCM-D measurements.
Results AFM Observation of Polymeric Bilayers. Figure 2 shows in situ AFM images of a patterned polymeric bilayer. The gridshaped area in Figure 2A is the polymeric bilayer (UV irradiation: 5.0 J/cm2), whereas the square-shaped areas are voids formed after removing monomers by SDS. Figure 2B is a magnified view of the boundary region marked in (A). The boundaries of polymeric bilayers were rather uneven, presumably due to the limited resolution of the currently applied photolithography setup. The bilayer thickness was 5.1 nm, as shown in the line cross section (Figure 2C). The value agrees well with the previous study using contact mode AFM in water (4.6 ( 0.5 nm).21,22 It should be noted that the thickness obtained in the present study should include the water layer between the substrate and the bilayer, since we worked with the tapping mode. We studied the morphologies of polymerized bilayers as a function of applied UV irradiation doses. Representative images are shown in Figure 3. The number of remaining polymeric bilayer domains increased with the UV dose. In the case of the smallest UV irradiation dose (1.5 J/cm2), oval bilayer disks were randomly distributed on the glass surface (Figure 3A). The thickness of disks was ∼5 nm, consistent with the value expected for bilayers. It is interesting to note that we seldom observed monolayer domains, although the polymerization should occur independently within each monolayer.17 This result may imply that monolayers of polymerized DiynePC are less stable toward the SDS treatment compared with bilayers.21 The size distribution of polymeric bilayer domains is shown in Figure 4 as a histogram of the radius (we calculated from the measured areas assuming a circular shape). The average radius of the bilayer domains was ∼13 nm. This value, however, is a slight overestimation, because AFM probes have a finite curvature at their tip (Vide infra). (21) Morigaki, K.; Scho¨nherr, H.; Frank, C. W.; Knoll, W. Langmuir 2003, 19, 6994–7002. (22) Morigaki, K.; Scho¨nherr, H.; Okazaki, T. Langmuir 2007, 23, 12254– 12260.
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Figure 3. AFM images (1 × 1 µm2) of polymeric bilayers with varied UV application doses: (A) 1.5 J/cm2, (B) 2.5 J/cm2, and (C) 4.0 J/cm2.
Figure 4. Size distribution of the polymeric bilayer domains observed after the SDS treatment (the UV dose was 1.5 J/cm2). The radius of domains was plotted in a histogram, assuming a circular shape. The influence of finite AFM probe curvatures is not corrected.
Figure 3B shows the sample with a UV irradiation dose of 2.5 J/cm2. In this image, the surface appears to be mostly covered by the polymer, with some pinholelike defects (pits). The depth of the larger pits was measured to be consistent with the bilayer thickness, whereas smaller pits could not be measured due to the lateral resolution of the AFM probe in this experimental condition (data not shown). For the applied UV dose of 4.0 J/cm2, the surface was mostly covered with the polymer and some protrusions were observed (Figure 3C). The protrusions had multiple terraces, with the height ranging from 6 to 30 nm. They were presumably bilayers ejected from the DiynePC SPB due to its expansion in the lateral direction upon polymerization. It has been reported that diacetylene phospholipid bilayers expand in the lateral direction and shrink in the vertical direction during polymerization because of the orientation changes of the diacetylene moiety (see the schematic illustration shown in Figure 1E).23 The structural changes occurring in the polymerization were confirmed also by directly comparing monomeric and polymeric bilayers (Figure 5). A patterned polymeric bilayer (UV dose: 5 J/cm2) was observed by AFM without removing monomers by the SDS treatment. The surface of the monomeric bilayer was ∼1.6 nm higher than that of the polymeric bilayer (Figure 5C), suggesting bilayer shrinkage in the vertical direction upon polymerization. The magnified image (Figure 5B) shows that there are many pits in the monomeric bilayer region (left part), while the pits mostly disappeared and there are some protrusions in the polymeric bilayer (right part). Pits in the monomeric bilayers should be formed in the process of assembling the bilayer by the LB/LS technique. Our previous AFM studies suggested that the LS deposition was mainly responsible for the (23) Bourdieu, L.; Chatenay, D.; Daillant, J.; Luzet, D. J. Phys. II 1994, 4, 37–58.
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Figure 5. (A) AFM image (20 × 20 µm2) of a patterned polymeric bilayer without the SDS treatment, where both monomeric and polymeric bilayers were observed. (B) Magnified image (3.5 × 3.5 µm2) of the boundary marked in (A). (C) Line profile measured along the line in (B) for the determination of the height difference.
generation of pits, because the first monolayer had a much smaller number of defects.21 The reduced number of pits in the polymeric bilayers strongly suggests that the polymeric bilayers expanded laterally, filling the pits and inducing some protrusions due to the collapse of bilayers. Determination of the Polymeric-Fluid Lipid Compositions. Lipid membranes can be incorporated into the voids of partially polymerized bilayers, forming a composite membrane of polymeric and fluid bilayers. Since the amount of polymerized bilayer depends on the applied UV irradiation dose, as shown in Figure 3, the composition of polymeric and fluid bilayers can be modulated by changing the UV dose. The amount of polymerized and fluid bilayers was quantified as a function of the UV irradiation dose. Two independent methods, ellipsometry and fluorescence microscopy, were applied for the estimation. We did not use AFM to estimate the coverage by polymeric bilayers, because AFM probes cannot identify smaller defects below their tip diameters (ca. 40 nm) and it should overestimate the area fraction (Vide infra). We also did not use fluorescence from polymerized DiynePC for the estimation, since the fluorescence intensity does not correlate linearly with the amount of polymeric bilayers.15 Figure 6A shows the amount of polymeric bilayers estimated by ellipsometry. The amount of polymeric bilayers is given as an average thickness. Since the thickness of individual bilayer domains should be constant, as shown by AFM observation, a higher thickness indicates a higher coverage by polymeric bilayers. It should be noted that bilayer samples were prepared on silicon wafers covered with a native oxide layer for these measurements. Although silicon wafers and glass slides are significantly different materials, we assume that they provide similar conditions for the polymerization of DiynePC, because the surface properties and roughness are very similar between them. Furthermore, although silicon wafers are reflecting light, the effect of interference between the incident and reflected lights would be negligible, because the oxide layer is very thin (ca. 1 nm). The average thickness increased as a function of the applied UV irradiation dose, reaching a plateau between 4.0 and 5.5 J/cm2, and slightly decreasing above 6.0 J/cm2. The amount of incorporated fluid bilayers within the matrix of partially polymerized DiynePC bilayers was estimated by fluorescence microscopy. We prepared composite membranes of polymeric DiynePC and fluid bilayers (egg-PC containing 1 mol % TR-PE) and measured fluorescence arising from the fluid bilayer fractions (see the Supporting Information). We varied the UV irradiation doses (1.5-5.0 J/cm2), and the fluorescence intensity was plotted versus the amount of polymerized DiynePC bilayers determined by ellipsometry (Figure 6B). The amount of incorporated lipids decreased linearly with the amount of
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Figure 7. Changes of the resonant frequency (∆f) and dissipation (∆D) measured by QCM-D during vesicle fusion on SiO2 (black) and polymerized DiynePC surfaces. The UV irradiation dose for the photopolymerization was 4.0 J/cm2 (light blue), 2.0 J/cm2 (blue), 1.0 J/cm2 (green), and 0.5 J/cm2 (red).
Figure 6. (A) Amount of polymerized DiynePC that remained on the substrate after the SDS treatment as measured by ellipsometry and plotted versus UV irradiation dose. The thickness represents an average value of polymeric bilayers and voids. (B) Fluorescence intensity of TR-PE arising from incorporated lipid bilayers plotted versus the amount of polymerized DiynePC measured by ellipsometry.
preformed polymeric bilayers. It is a clear indication that polymeric and fluid bilayers are forming a single layer of a composite membrane. The linearity also demonstrates the consistency between the two evaluation methods (ellipsometry and fluorescence). Interestingly, the fluorescence intensity from TR-PE was not zero for samples with the maximum polymer coverage. It may suggest that polymeric bilayers do not completely cover the surface even for the maximum coverage reached in the present experimental conditions. An alternative possibility is that there are some adsorbed lipid materials on the polymer surfaces. However, based on the QCM-D results, we do not suppose that this factor has a major contribution to the observed fluorescence (Vide infra). By extrapolating the linear relation to the x-axis, the theoretical thickness for complete bilayer coverage could be estimated to be 4.4 nm. QCM-D Monitoring of the Bilayer Incorporation. In addition to determining the lipid compositions, the structural integrity of polymeric and fluid bilayers was studied by the QCM-D technique. We prepared polymeric DiynePC bilayers on sensor plates with different UV doses and monitored the frequency and dissipation shifts (∆f and ∆D) upon incorporation of fluid bilayers (140 µM egg-PC vesicles containing 1 mol % TR-PE).24 The measured changes in ∆f and ∆D (fifth harmonics: 25 MHz) are shown in Figure 7. On an SiO2 substrate without polymeric bilayers, a characteristic two-phase process with a hump was observed due to the accumulation of vesicles before the formation of SPBs (black line).25 The final ∆f value of ∼26 Hz is consistent for the adsorption of single SPB on the (24) It should be noted that the UV doses in these experiments are not identical to those in Figure 7 due to a rather thick SiO2 layer of the sensor chips (ca. 50 nm), which causes interference between incident and reflected light. (25) Keller, C. A.; Kasemo, B. Biophys. J. 1998, 75, 1397–1402.
surface.25-27 On the other hand, no or little humps of ∆f and ∆D were observed for the substrates with polymeric bilayers (red and green lines). The absence of humps should indicate a rapid transformation of adsorbed vesicles into SPB on the surface. From our previous studies, the accelerated SPB formation should be due to the effect of polymeric bilayer edges that destabilize adsorbed vesicles.19 Partially polymerized bilayers composed of small domains should have a high density of bilayer edges. One interesting observation is the increase in ∆f and decrease in ∆D observed in the sample with the 2.0 J/cm2 UV irradiation dose (blue line), which is opposite to normally observed behaviors. This may be due to the effect of water molecules that were trapped in the gaps between partially polymerized bilayers. It is plausible that adsorption of lipid bilayers in the gaps released trapped water molecules. The net effect on ∆f and ∆D should depend on the amount of water molecule desorption and lipid bilayer adsorption. Neither ∆f nor ∆D changed on the substrates that had a maximum amount of polymerized bilayer (4.0 J/cm2) (light blue line). Obstruction of Lateral Diffusion by Polymeric Bilayer Domains. Polymeric bilayer domains can act as an obstacle for the lateral diffusion of membrane-associated molecules. We determined the degree of obstruction as a function of the fraction of polymeric bilayers. The lateral diffusion coefficients of lipids (TR-PE) were measured by using the FRAP method with the BPE analyses. In the case of egg-PC/TR-PE bilayers on a glass substrate (no polymer), the average diffusion coefficient was determined to be 1.6 ( 0.4 µm2/s. This value agrees well with the results in previous reports.20,28-31 The results of partially polymerized DiynePC bilayers are compiled in Figure 8. The diffusion coefficients were normalized to the obstacle-free diffusion (i.e., 1.6 µm2/s) and expressed as relative diffusion coefficients, D*. The area fractions of polymeric bilayers, c, on the other hand, were derived by dividing the film thickness from ellipsometric measurements with the theoretical thickness cor47.
(26) Richter, R.; Mukhopadhyay, A.; Brisson, A. Biophys. J. 2003, 85, 3035–
(27) Reimhult, E.; Ho¨o¨k, F.; Kasemo, B. Langmuir 2003, 19, 1681–1691. (28) Chan, P. Y.; Lawrence, M. B.; Dustin, M. L.; Ferguson, L. M.; Golan, D. E.; Springer, T. A. J. Cell Biol. 1991, 115, 245–255. (29) Stelzle, M.; Miehlich, R.; Sackmann, E. Biophys. J. 1992, 63, 1346–1354. (30) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400–1414. (31) Deverall, M. A.; Gindl, E.; Sinner, E. K.; Besir, H.; Ruehe, J.; Saxton, M. J.; Naumann, C. A. Biophys. J. 2005, 88, 1875–1886.
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Figure 8. Relative diffusion coefficients of TR-PE in the composite membranes plotted as a function of the area fraction of polymeric bilayers (obstacles) obtained from ellipsometry and fluorescence microscopy.
responding to the full coverage (i.e., 4.4 nm). The diffusion coefficients decreased monotonically as a function of the area fraction and were found to be nearly zero for a higher c (above ca. 0.7). The decrease was linear at low obstacle area fractions below 0.4. However, we observed that a finite lateral diffusion remained for the area fraction between 0.4 and 0.7, and we did not find a clear threshold at which the diffusion coefficient became zero.
Discussion We evaluated the morphology of polymerized DiynePC bilayer domains and their effects on the lateral mobility of lipids in integrated fluid bilayers. The AFM observations revealed that polymeric bilayer domains were smaller (an apparent mean radius of 13 nm) and more uniform in size compared with those observed in the previous study (compare Figure 11 ref 21 and Figure 3 in this paper). The difference in observed features can be attributed to the preparation conditions of monomeric bilayers. Diacetylene moieties undergo polymerization in the solid state,17,18 and the range of polymerization is limited to the domain of monomeric bilayers with crystalline packing. We have recently found that the domain sizes can be controlled by the preparation temperature at which a monolayer of DiynePC monomers is formed at the air/water interface.22 If a monolayer was formed below the triple point temperature of DiynePC monolayers (ca. 20 °C), a direct transition of DiynePC lipids from the gaseous state to the liquid condensed state occurred.32 It resulted in small domains of DiynePC in the monolayer, which could be transferred onto substrates by the LB/LS techniques. Samples prepared above the triple point temperature had larger domain sizes in the Langmuir monolayer and resulted in larger and fractal polymeric domains.21 The present AFM observation has also revealed the structural changes of DiynePC bilayers upon polymerization. The bilayer thickness decreased upon polymerization, as shown in Figure 5, and pinholelike defects observed in monomers disappeared in polymeric bilayers (Figure 5B) due to a lateral expansion of the film. These results are consistent with the previously reported behaviors of DiynePC polymerization in monolayers.23 The composition of polymerized and fluid bilayers could be modulated by changing the UV irradiation dose in the photopolymerization. The amount of residual polymer film after the SDS treatment did not increase linearly with the UV dose (Figure 6A). It is presumably due to the critical size of domains necessary for withstanding solubilization by the detergent. The amount of polymerized and fluid bilayers determined by ellipsometry and fluorescence microscopy, respectively, changed linearly (Figure 6B). This result strongly supports the premise that polymerized (32) Britt, D. W.; Hofmann, U. G.; Mo¨bius, D.; Hell, S. W. Langmuir 2001, 17, 3757–3765.
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and fluid bilayers are forming a single layer of a composite membrane. One important finding is the possibility that polymeric bilayers do not cover the entire surface even for the maximum coverage achieved in the present experimental conditions. Minute defects in the form of either pinholes or narrow cracks seemed to remain. The exact value of maximum polymer coverage could not be determined due to the possibility that adsorbed vesicles on the surface of polymeric bilayers were influencing the estimation, although the QCM-D results suggested that the amount of adsorbed vesicles should be rather small. From the AFM observations, we suppose that the increase of polymer coverage was due to the increase in the number of polymeric bilayer domains without significant changes in their sizes. This conclusion should be reasonable from the fact that sizes of polymeric bilayer domains are mainly determined by the sizes of monomeric bilayer domains. It was supported also by comparing the polymer area fractions obtained by fluorescence microscopy and AFM. AFM measurements tended to overestimate the coverage due to the curvature of the tip. The area fractions of polymeric bilayers estimated by ellipsometry and AFM were proportional, with the AFM results being roughly twice those of fluorescence microscopy (see the Supporting Information). The overestimation by AFM should be size-dependent, with small domains being more strongly affected. The fact that the ratio between the areas determined by the two methods remained constant should be a strong indication that there was not a significant change in the domain size distribution. From the proportion of ∼2 between AFM and ellipsometry, we could estimate the actual mean size of the polymeric bilayer domains to be 9.5 nm, assuming again a circular shape for the domains. The effect of polymeric DiynePC domains on long-range lateral diffusion of mobile lipids was quantified by FRAP measurements. The diffusion coefficients decreased monotonically with the amount of polymeric bilayer domains and reached zero for a higher c (above ca. 0.7). The decrease was linear at low obstacle area fractions below 0.4. The obstruction of lateral diffusion in two-dimensional fluid systems has been extensively studied by using the percolation model.31,33-44 The effect of obstruction depends on various factors such as the size of obstacles and the interactions between mobile species and obstacles. It has been observed that smaller obstacles are more effective in obstructing lateral diffusion.33,44,45 The effective obstruction observed in the linear regime (below 0.4) should be ascribed to the small size of the polymeric bilayer domains. Another important factor is the effect of polymeric bilayers on fluid bilayers at their boundaries. We assessed this by analyzing the data with a modified free-area model, which incorporates a soft-core repulsion between obstacles and diffusing molecules due to the presence of ordered lipid molecules at the boundary.39 The solid line in Figure 8 is the least-squares fitting result using the following second-order polynomial and coefficients (a and b). (33) Saxton, M. J. Curr. Top. Membr. 1999, 48, 229–282. (34) Saxton, M. J. Biophys. J. 1989, 56, 615–622. (35) Vaz, W. L. C.; Melo, E. C. C.; Thompson, T. E. Biophys. J. 1989, 56, 869–876. (36) Bultmann, T.; Vaz, W. L. C.; Melo, E. C. C.; Sisk, R. B.; Thompson, T. E. Biochemistry 1991, 30, 5573–5579. (37) Saxton, M. J. Biophys. J. 1992, 61, 119–128. (38) Saxton, M. J. Biophys. J. 1993, 64, 1053–1062. (39) Almeida, P. F. F.; Vaz, W. L. C.; Thompson, T. E. Biochemistry 1992, 31, 7198–7210. (40) Saxton, M. J. Biophys. J. 1994, 66, 394–401. (41) Schram, V.; Tocanne, J. F.; Lopez, A. Eur. Biophys. J. 1994, 23, 337– 348. (42) Schram, V.; Lin, H. N.; Thompson, T. E. Biophys. J. 1996, 71, 1811– 1822. (43) Saxton, M. J. Biophys. J. 1997, 72, 1744–1753. (44) Ratto, T. V.; Longo, M. L. Biophys. J. 2002, 83, 3380–3392. (45) Eisinger, J.; Flores, J.; Petersen, W. P. Biophys. J. 1986, 49, 987–1001.
Polymerized Lipid Bilayers on a Solid Substrate
D/ ) 1 + ac + bc2 a ) -1.208 - 24.3exp(-1.763R ⁄ ξ) 2.408exp(-0.3138R ⁄ ξ) b ) 185exp(-2.587R ⁄ ξ) R is the radius of the obstacles, and ξ is the coherence length of the obstacles, which represents the degree of ordering in mobile lipids at the boundaries to the obstacles. We obtained the value of R/ξ ) 2.12 by fitting the result. By using the average radius of polymerized bilayer domain obtained from the AFM measurements and corrected by taking the ellipsometry results into account (9.5 nm), the coherence length was calculated to be 4.5 nm. This value is considerably larger than the theoretical prediction of ∼2.5 nm from thermodynamic considerations,46 but it fits within the range reported previously for mixed bilayers composed of solid-phase and fluid-phase phospholipid bilayer domains (4.5-6.5 nm).44 This result indicates that polymeric DiynePC domains affect the ordering of mobile lipids at the boundaries in the same manner as solid-phase phospholipid domains. As the polymeric DiynePC coverage is increased, the obstacles should connect each other and separate the continuous conducting phase at the percolation threshold. However, in the present study, D* did not reach zero at the extrapolation of the slope below 0.4. At the area fraction between 0.4 and 0.7, a finite diffusion persisted. The less effective obstruction in this regime may be ascribed to the presence of channel-like defects in polymeric bilayers. Although we could not directly observe the channels by AFM, the discrepancy between the areas measured by AFM and fluorescence measurements is a strong indication for the presence of these channels that cannot be detected by an AFM probe. The channel-like defects between the domains should be formed by the mechanical stress of the polymerization with constraints in the film curvature and occupied area.23,32 They seem to persist up to a relatively high coverage of polymeric bilayers. Practically no long-range lateral diffusion was observed for the polymeric coverage above 0.7. (46) Ja¨hnig, F. Biophys. J. 1981, 36, 329–345.
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In summary, we obtained the following quantitative information on the structure and physicochemical properties of polymeric lipid bilayers on a solid substrate. First, AFM observations revealed that polymerized DiynePC bilayers were formed as nanometric domains of a relatively well-defined size distribution. Second, quantification of polymeric and fluid bilayer fractions by ellipsometry and fluorescence microscopy demonstrated that these membranes are integrated as a single layer of a composite membrane with defined ratios. Third, the polymeric bilayers were effective in obstructing the long-range diffusion of lipids, and the degree of obstruction could be quantitatively modulated by the amount of polymeric bilayer domains. These findings should be of great importance for designing model membrane systems with well-defined structures and physicochemical properties. The compositions of polymeric and fluid bilayer fractions can be spatially controlled by lithographically changing the UV irradiation dose. Therefore, by designing the geometry and degree of polymerization, it should be possible to utilize composite membranes in biophysical studies such as mimicking microdomains of the biological membranes as well as various biomedical applications including electrophoretic separation of membrane components. Acknowledgment. We thank Ms. Saori Mori and Ms. Maki Koike for their assistance in the preparation of patterned substrates, vesicle suspensions, ellipsometric measurements, and fluorescence microscopy observations. We thank Dr. Junji Nishii and Dr. Kenji Kintaka (Photonics Research Institute, AIST) for allowing us to use the ellipsometer. This work has been supported in part by Promotion Budget for Science and Technology (AIST Upbringing of Talent in Nanobiotechnology Course) from the Ministry of Education, Science, Culture and Sports (MEXT) and by the Joint Studies Program of the Institute for Molecular Science. Supporting Information Available: Method used for estimating the amount of incorporated fluid lipid bilayers by fluorescence microscopy (Figure S1) and comparison of the area fractions of polymeric bilayers estimated by ellipsometry and AFM (Figure S2). This material is available free of charge via the Internet at http://pubs.acs.org. LA802670T
