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Polymerization of Diacetylene Phospholipid Bilayers on Solid Substrate: Influence of the Film Deposition Temperature Kenichi Morigaki,*,† Holger Scho¨nherr,*,‡ and Takashi Okazaki†,§ Research Institute for Cell Engineering, National Institute of AdVanced Industrial Science and Technology (AIST), Ikeda 563-8577, Japan, UniVersity of Twente, MESA+ Institute for Nanotechnology and Faculty of Science and Technology, Department of Materials Science and Technology of Polymers, P.O. Box 217, 7500 AE Enschede, The Netherlands, Graduate School of Science, Osaka UniVersity, Toyonaka 560-0043, Japan ReceiVed May 9, 2007. In Final Form: August 4, 2007 Micropatterned phospholipid bilayers on solid substrates offer an attractive platform for various applications, such as high throughput drug screening. We have previously developed a photopolymerization-based methodology for generating micropatterned bilayers composed of polymerized and fluid lipid bilayers. Lithographic photopolymerization of a diacetylene-containing phospholipid (DiynePC) allowed facile fabrication of compartmentalized arrays of fluid lipid membranes. Herein, we report on a key experimental parameter that significantly influences the homogeneity and quality of the fabricated polymeric bilayers, namely the temperature at which monolayers of monomeric DiynePC were formed on the water surface and transferred onto solid substrates by the Langmuir-Blodgett/Langmuir-Schaefer (LB/LS) technique. Using fluorescence microscopy and atomic force microscopy, it was found that polymerized bilayers were homogeneous, if bilayers of DiynePC were prepared below the triple point temperature (ca. 20 °C) of the monolayer, where a direct transition from the gaseous state to the liquid condensed state occurred. Bilayers prepared above this temperature had a markedly increased number of crack-like line defects. The differences were attributed to the domain structures in the monolayer that were transferred from the water surface to the substrate. Domain size, rather than the molecular packing in each domain, was concluded to play a critical role in the formation of defects. The spontaneous curvature and area changes of bilayers were postulated to cause destabilization and detachment of the films from the substrate upon polymerization. Our present results highlight the importance of controlling the domain structures for the homogeneity of polymerized bilayers required in technological applications.

Introduction Substrate-supported planar lipid bilayers (abbreviated as SPBs in the following) provide unique possibilities for reconstituting cellular membranes on solid surfaces.1-3 Micropatterning of SPBs represents in this context an attractive approach that allows the creation of designed arrays of biological materials and should facilitate various new applications, such as high throughput drug screening.4-6 One pertinent shortcoming of SPBs is their inherent limited stability. Possible pathways to improve the stability include the use of self-assembled monolayers,7 tethered lipo-polymers to the surface,8-10 and polymerizable lipids.11-13 * Corresponding authors. Tel.: ++81-72-751-4142. Fax: ++81-72751-9628. E-mail: [email protected]. Tel.: ++31 53 489 3170. Fax ++31 53 489 3823. E-mail: [email protected]. † AIST. ‡ University of Twente. § Osaka University. (1) Sackmann, E. Science (Washington) 1996, 271, 43-48. (2) Tanaka, M.; Sackmann, E. Nature 2005, 437, 656-663. (3) Scho¨nherr, H.; Degenhart, G. H.; Dordi, B.; Feng, C. L.; Rozkiewicz, D. I.; Shovsky, A.; Vancso, G. J. AdV. Polym. Sci. 2006, 200, 169-208. (4) Groves, J. T.; Ulman, N.; Boxer, S. G. Science (Washington) 1997, 275, 651-653. (5) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149-157. (6) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124, 23942395. (7) Plant, A. Langmuir 1999, 15, 5128-5135. (8) Spinke, J.; Yang, J.; Wolf, H.; Liley, M.; Ringsdorf, H.; Knoll, W. Biophys. J. 1992, 63, 1667-1671. (9) Schiller, S. M.; Naumann, R.; Lovejoy, K.; Kunz, H.; Knoll, W. Angew. Chem., Int. Ed. 2003, 42, 208-211. (10) Atanasov, V.; Knorr, N.; Duran, R. S.; Ingebrandt, S.; Offenha¨usser, A.; Knoll, W.; Ko¨per, I. Biophys. J. 2005, 89, 1780-1788. (11) Ringsdorf, H.; Schlarb, B.; Venzmer, J. Angew. Chem., Int. Ed. 1988, 27, 113-158. (12) Mueller, A.; O’Brien, D. F. Chem. ReV. 2002, 102, 727-757.

To realize stable SPBs, we have recently developed a methodology to create micropatterned SPBs composed of polymerized and fluid lipid bilayers, where the polymerized bilayer forms an integrated matrix with embedded fluid bilayer corrals (Figure 1).14-17 For the polymerization of bilayers, a commercially available diacetylene phospholipid, 1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine (DiynePC), has been used (Figure 1). After the UV polymerization, monomers were removed by subsequent treatment with a detergent solution, and various phospholipid bilayers could be incorporated into the lipid-free regions by the vesicle fusion method, forming a hybrid membrane composed of polymeric and fluid lipid bilayers. In this configuration, polymeric bilayers act both as a barrier for the lateral diffusion of membrane-associated molecules and as a stabilizing element for the SPBs. Therefore, the generation of polymeric bilayers in a controlled manner is critical for this micopatterning strategy. It is generally well-established that diacetylenes polymerize in the solid state (topochemical polymerization) and that the polymerization efficiency depends very sensitively on the molecular packing of the diacetylene monomers.18-20 We have (13) Ross, E. E.; Bondurant, B.; Spratt, T.; Conboy, J. C.; O’Brien, D. F.; Saavedra, S. S. Langmuir 2001, 17, 2305-2307. (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.; Scho¨nherr, H.; Frank, C. W.; Knoll, W. Langmuir 2003, 19, 6994-7002. (17) Morigaki, K.; Kiyosue, K.; Taguchi, T. Langmuir 2004, 20, 7729-7735. (18) Wegner, G. Makromol. Chem. 1972, 154, 35-48. (19) Bloor, D.; Chance, R. R. Polydiacetylenes: Synthesis, Structure and Electronic Properties; Martinus Nijhoff Publishers: Dordrecht, The Netherlands, 1985.

10.1021/la701346x CCC: $37.00 © 2007 American Chemical Society Published on Web 10/18/2007

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Materials and Methods

Figure 1. (A) Schematic of the pattering method, and (B) the structure and polymerization scheme of DiynePC.

previously found that successful cross-linking of bilayers depended on various factors such as the structure of the amphiphilic molecules and the substrate material.15,21 Under certain conditions, polymerization proceeded within the SPB while the bilayer structure is retained, as revealed by the atomic force microscopy (AFM) observations.16 In the course of our investigations, we realized that the morphologies of polymerized DiynePC bilayers were drastically affected by the temperature at which the monolayers were formed on the water surface and transferred onto solid substrates by the LB/LS technique. Due to the importance of fabricating defect-free polymeric bilayers for defining the corrals and preventing unwanted mixing of incorporated fluid lipid bilayers, the relation between deposition temperature and bilayer morphologies has thus been studied. In the present paper, we report on this temperature effect in detail, providing insights into the polymerization mechanism of thin films and practical information for the fabrication of micropatterned bilayers on solid substrates. (20) Cantow, H.-J. Polydiacetylenes; Springer-Verlag: Berlin, 1984. (21) Each DiynePC molecule has two diacetylene moieties that can be polymerized independently and they are not incorporated in the same polymer chain (Lopez, E.; O’Brien, D. F.; Whitesides, T. H. J. Am. Chem. Soc. 1982, 104, 305-307). The polymerization proceeds predominantly within the same monolayer. Therefore, two layers of cross-linked networks are formed within bilayers upon photopolymerization.

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-phosphoethanolamine (TR-PE) was purchased from Molecular Probes (Eugene, OR). Sodium dodecyl sulfate (SDS) was purchased from Nacalai Tesque (Kyoto, Japan). All chemicals were reagent grade and used without further purification. The 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 the substrates, for preparing the buffer solution (0.01 M phosphate buffer with 0.15 M NaCl, pH 6.6), and as the imaging medium for AFM analyses. Substrate Cleaning. Microscopy glass slides (Matsunami, Osaka, Japan) were used as substrates. 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 0.05:1:5 NH4OH (28%): H2O2 (30%):H2O for 10 min at 65 °C, rinsed again with deionized water extensively, and then dried in a vacuum oven for 30 min at 80 °C. This protocol resulted in clean and hydrophilic surfaces for the adsorption of lipid bilayer membranes. Preparation of Patterned DiynePC Bilayers. Polymerizable 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 (HBM-AP, Kyowa Interface Science, Asaka, Japan). The temperature of the subphase (deionized water) was controlled by circulating thermostated water beneath the trough. 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 the 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 was conducted by UV irradiation using a mercury lamp (UVE-502SD, Ushio, Tokyo, Japan) as the light source. The polymerization was conducted in a closed system that was comprised of a deionized water reservoir, a pump, and a cell (∼4 mL volume). The water reservoir was depleted of oxygen by purging it with argon. Oxygen-free water was circulated continuously by the pump through the cell where polymerization of the bilayers was conducted. The cell had two walls on opposite sides, 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. Desired patterns were transferred to the DiynePC bilayer in the polymerization process by illuminating the sample through a mask (a quartz slide with a patterned chromium layer coating) which was placed directly on the DiynePC bilayer. 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 irradiation dose was 4 J/cm2, which was previously shown to be sufficient to form a cross-linked polymeric DiynePC bilayer.17 After the UV irradiation, nonpolymerized DiynePC molecules were removed from the substrate surface by treatment in 0.1 M SDS solution at 30 °C for 30 min, and extensive rinsing with deionized water. The patterned polymeric DiynePC substrates were stored in deionized water in the dark at 4 °C. Fluorescence Microscopy. Fluorescence microscopy was conducted 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 bilayers were observed by the Olympus U-MNIBA2 filter set (excitation wave-

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Figure 3. Fluorescence micrographs of polymeric bilayers prepared by the LB/LS method at 16 and 28 °C, followed by SDS treatment and backfilling with egg-PC. The polymeric bilayers were observed by the NIBA filter set (left side), whereas the backfilled fluid membranes were selectively observed by the WIY filter set (right side). The scale bar corresponds to 20 µm. Figure 2. π-A isotherms of DiynePC Langmuir monolayers at 16 and 28 °C. length: 470-490 nm, emission wavelength: 510-550 nm: abbreviated as NIBA), and TR-PE was observed by the Olympus U-MWIY2 filter set (excitation wavelength: 545-580 nm, emission wavelength: >610 nm: abbreviated as WIY). Fluorescence images were collected with a CCD camera (DP30BW, Olympus) and processed with the MetaMorph program (Molecular Devices, Sunnyvale, CA). Atomic Force Microscopy (AFM). The AFM data were acquired on a VEECO/Digital Instruments (DI) multimode AFM equipped with a NanoScope IIIa controller (DI, Santa Barbara, CA) in contact mode (CM) in water using a DI liquid cell. V-shaped or single beam Si3N4 cantilevers with various spring constants were used (Model NP, Veeco Nano Probe, Santa Barbara, CA knominal ) 0.06-0.58 N/m and model OMCL-RC800PSA, Olympus, Tokyo, Japan, knominal ) 0.02 N/m, respectively). The scanner was protected against liquid spillage by covering the top part with a very thin film of sealant (Parafilm, American National Can, Neenah, WI). Samples were mounted into the AFM while keeping the surface, which was later imaged, covered by water at all times, as described previously.16 All of the images shown here were subjected to a first order planefitting procedure to compensate for sample tilt and, if necessary, to a zerothorder flattening.

Results We found that the morphology of polymerized bilayers was altered drastically depending on whether monomeric bilayers were assembled below or above the triple point temperature of DiynePC monolayer on the water surface (20 °C, vide infra). Therefore, we transferred monolayers of DiynePC monomer from the air/water interface to glass substrates either at 16 °C or at 28 °C and compared the morphologies of bilayers microscopically after polymerization and SDS treatment. Pressure-Area (π-A) Isotherms of the Monolayer. Figure 2 compares the π-A isotherms of DiynePC on water surface at 16 and 28 °C. The isotherm at 16 °C showed a direct transition from the gas phase to the liquid condensed phase, in accordance with previous observations (vide infra), whereas the isotherm at 28 °C went through the liquid expanded phase before reaching the liquid condensed phase.22,23 An “overshoot” at the transition between liquid expanded-liquid condensed phases was observed for the isotherm at 28 °C. The overshoot hump was observed only in the first compression. A plateau region was observed for the liquid expanded-liquid condensed transition in later com(22) Johnston, D. S.; McLean, L. R.; Whittam, M. A.; Clark, A. D.; Chapman, D. Biochemistry 1983, 22, 3194-3202. (23) Bourdieu, L.; Chatenay, D.; Daillant, J.; Luzet, D. J. Phys. II 1994, 4, 37-58.

pressions (data not shown), in accordance with previous observations with other diacetylene phosphatidylcholine monolayers.24 Bourdieu et al. reported that 20 °C was the triple point temperature of DiynePC monolayer on water surface.23 Below this temperature, a direct transition from a gaseous state to a liquid condensed state occurs, whereas a coexistence plateau between the liquid expanded and liquid condensed phases is observed above this temperature. Our data are in full agreement with this report. Fluorescence Microscopic Observations. Figure 3 shows fluorescence micrographs of lithographically polymerized DiynePC bilayers that were deposited from the water surface either at 16 or at 28 °C. Except for the film deposition temperatures, all subsequent procedures for the bilayer polymerization, monomers removal, and backfilling with fluid phospholipid bilayers (eggPC/TR-PE) were exactly the same for both samples. The polymeric bilayer was homogeneous to the resolution of fluorescence microscopy and mostly void of fluid membranes, if the monomers were deposited at 16 °C. On the other hand, the polymeric bilayer deposited at 28 °C showed a markedly increased number of crack-like line defects that were also visualized with incorporated bilayers of egg-PC/TR-PE. Since UV polymerization was conducted at the same temperature for both samples, the observed differences in morphologies should stem from the difference in the film deposition temperature. The defects were observed also before the SDS treatment, indicating that they were formed upon bilayer polymerization.25 Effect of Annealing Monomeric Bilayers. The observed difference of bilayer morphologies in Figure 3 points to the influence of temperature at which monomeric bilayers were formed. Therefore, we studied the effects of annealing by incubating monomeric bilayers at an elevated temperature, 60 °C, and cooling down to 10 °C. Since the phase transition temperature of DiynePC bilayer was reported to be 38 °C,22 bilayers should be in the liquid crystalline state at 60 °C. The fluorescence micrographs of polymeric bilayers are shown in Figure 4. For both samples prepared at 16 and 28 °C, the annealing process significantly altered the morphologies of polymerized bilayers and mostly eliminated the differences due to the film deposition temperatures. Importantly, we found that the morphologies of polymeric bilayers were strongly affected by the cooling rate. For the samples that were cooled rapidly (the sample was immersed into a water bath of 10 °C), polymeric bilayers (24) Hui, S. W.; Yu, H.; Xu, Z.; Bittman, R. Langmuir 1992, 8, 2724-2729. (25) We have previously observed the morphology of monomers and bilayers composed of monomeric DiynePC at 25 °C (ref 16). The films were mostly void of defects, except for some small pin-holes. Therefore, we suppose that the cracklike defects were not formed upon bilayer deposition.

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of high and low-temperature samples were also identical to within the experimental error. The values of the layer thickness (analysis of single line cross-section after plane-fit, arithmetic mean ( standard deviation) were (4.8 ( 0.4) nm for samples prepared at 28 °C and (4.6 ( 0.5) nm for films deposited at 16 °C. However, a small but significant difference in rms roughness was detected for the two series of samples (Figure 6): samples prepared at high temperature showed an increased rms roughness. Finally, AFM provided clear evidence for the successful removal of unpolymerized lipids in the bilayer film. The lower part of corrals was found to possess markedly different friction force26 and adherence properties27,28 in contact with the AFM tip as compared to the polymerized lipid bilayer (Figures 7 and 8). The friction forces experienced by the AFM tip on the polymerized lipid bilayer matrix were higher than in the corrals. The pull-off forces showed a consistent difference.

Discussion Figure 4. Fluorescence microscopy analysis of the effect of annealing on monomer film morphology: Monomeric bilayers deposited either at 16 or at 28 °C were thermally annealed before the polymerization by heating to 60 °C and subsequently cooling back to 10 °C. After the thermal treatments, the bilayers were polymerized, monomers were removed in 0.1 M SDS, and finally fluid bilayers (egg-PC/ TR-PE) were backfilled. The upper two rows show images captured on polymerized samples for which the monomer films were deposited at 16 °C. The lower two rows show images of corresponding films for which the monomer was deposited at 28 °C. The left, middle, and right columns show data of samples polymerized without annealing, samples cooled rapidly, and samples cooled slowly (in 2 h), respectively. The polymeric bilayers were observed by the NIBA filter set, whereas the backfilled fluid membranes were selectively observed by the WIY filter set. The size of corrals was 50 µm.

were generally homogeneous. Although we observed faint domain structures in the polymeric bilayers (indicated by arrows in Figure 4), whose presence was also visualized by a small amount of adsorbed egg-PC/TR-PE, the polymeric bilayers were mostly void of large-scale defects. Slow cooling over 2 h (rate ∼0.4 °C/min), on the other hand, resulted in heterogeneous polymeric bilayers with fractal domain structures and numerous defects. The polymeric bilayers also had significant amounts of egg-PC/TR-PE that were either adsorbed on the surface as vesicles or penetrating into the defects of polymeric bilayers. AFM Observations. The AFM observations provided complementary information on (1) homogeneity of the patterned layers and defect structures on optically inaccessible length scales, (2) surface roughness, (3) bilayer film thickness, and (4) surface properties of the samples prepared at different temperatures. Figure 5 displays contact mode AFM images of various scan sizes of patterned bilayers acquired in water. Clear differences in the defect morphology can be recognized among samples prepared at 28 °C (Figure 5, left column) and those prepared at 16 °C (Figure 5, right column). AFM height images of samples prepared at 28 °C displayed defects in the form of channels (up to ∼500 nm wide), whereas the samples prepared at 16 °C did not show such defects. These morphological differences are in agreement with the fluorescence microscopy data after backfilling with lipids, as shown in Figure 3. Higher resolution images displayed isolated point-like defects (which span ∼100-200 nm) for the latter samples. The hightemperature layers, by contrast, exposed likely the bare glass substrate in the mentioned channels, as well as in isolated pointlike defects (which span 100-200 nm, sometimes >700 nm). The depth of depressions assessed at the corral edges and at defect sites did not show significant differences. The step heights

We studied the influence of the deposition temperature of monomeric DiynePC on the bilayer morphology after polymerization. Polymeric DiynePC bilayers were observed to be homogeneous in the micrometer scale, if monomers were deposited from the water surface below the triple point temperature of DiynePC monolayer. AFM images obtained in situ provided evidence for the formation of nearly perfect corrals for bilayers fabricated at 16 °C (Figure 5). The contrast in AFM friction and pull-off force measurements is very pronounced and homogeneous (Figures 7 and 8), which implies that the bilayer deposition, polymerization and subsequent removal of monomeric lipid work well under the conditions employed. The experimentally determined bilayer thickness of ∼5 nm matches well with the expected bilayer thickness defined by the length of the lipids and the tilt angle of the molecules.16,23 By contrast, bilayers deposited above the triple point temperature showed a markedly increased number of crack-like line defects. Large scale defects were observed for samples prepared at 28 °C by fluorescence microscopy (e.g., in Figure 3, lower left panel) and AFM (Figure 5, panel a). The consequences of these defects are clear: the corrals are not enclosed and diffusive pathways to neighboring corrals may exist. Upon addition of additional vesicles, they will form a lipid bilayer outside the compartments (compare Figure 3). These defects potentially reduce the utility of micropatterned composite membranes by compromising the spatial definition given by the patterns. In addition, the edges of defects may result in the unwanted adsorption of vesicles.29,30 The rms roughness of the films determined by AFM is relatively small and does not yet approach the saturation for larger scan sizes, as expected. Interestingly, the roughness values were found to increase with the deposition temperature. In the plot of roughness vs square root of the scan size (Figure 6) this leads to an offset of approximately + 0.5 nm to larger values of the roughness. This observation is attributed to the increased number of defects in the high-temperature samples, which contribute to an increased roughness. We reason that the observed influence of the film deposition temperatures is closely related with the domain structures present (26) Tocha, E.; Scho¨nherr, H.; Vancso, G. J. Langmuir 2006, 22, 2340-2350. (27) Vancso, G. J.; Hillborg, H.; Scho¨nherr, H. AdV. Polym. Sci. 2005, 182, 55-129. (28) Scho¨nherr, H.; Vancso, G. J., Chemical Force Microscopy: Nanometer Scale Surface Analysis with Chemical Sensitivity. In SPM beyond imaging: Manipulation of Molecules and Nanostructures; Samori, P., Ed.; Wiley-VCH: Weinheim, Germany, 2006; pp 275-314. (29) Jenkins, A. T. A.; Bushby, R. J.; Evans, S. D.; Knoll, W.; Offenha¨usser, A.; Ogier, S. D. Langmuir 2002, 18, 3176-3180. (30) Scho¨nherr, H.; Rozkiewicz, D. I.; Vancso, G. J. Langmuir 2004, 20, 7308-7312.

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Figure 5. Contact mode AFM height images (acquired in water at RT) of a patterned bilayer for which monomer films were deposited at 28 (left) and at 16 °C (right). The samples were observed after the SDS treatment.

in the monolayer on water surface, and more specifically, with the size of the domains. As shown by the π-A isotherms (Figure 2), monolayers transformed directly from the gas phase into the liquid condensed phase at 16 °C, whereas they transformed via the liquid expanded phase at 28 °C. It is generally known that the courses of transitions affect the domain structures and sizes.31 It has been reported that a direct transition from the gas phase into the liquid condensed phase results in smaller domain sizes of monolayers compared with successive transitions from the gas phase into the liquid condensed phase via the liquid expanded phase.31 This can be partially explained by the kinetically hindered (31) Britt, D. W.; Hofmann, U. G.; Mobius, D.; Hell, S. W. Langmuir 2001, 17, 3757-3765.

nucleation of ordered patches of hydrocarbon chains and the concomitantly limited number of nucleation sites. The overshooting hump observed at 28 °C (Figure 2) also supports this interpretation. Although the overshooting hump was observed only in the first compression, we observed no morphological difference between polymeric bilayers prepared from monolayers deposited in the first compression and the second compression (data not shown). This result indicates that the microstructures of monolayers are determined not only by the initial nucleation but also by the route of phase transitions. Similar kinetic effects were observed also in the annealing experiments. Cooling down the monomeric bilayers rapidly from 60 to 10 °C generated more homogeneous bilayers after the polymerization, regardless of

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Figure 6. Values of rms roughness assessed in contact mode AFM height images (acquired in water at RT) of patterned bilayers after SDS treatment vs square root of AFM scan size. Monomer films were deposited either at 28 or at 16 °C.

the film deposition temperatures. These bilayers could effectively confine incorporated fluid bilayers to the imposed pattern. On the other hand, slow cooling over 2 h caused visible domain structures in the bilayers after the polymerization, and the number of defects and adsorbed vesicles in the polymeric bilayer also increased. Bilayers formed at different temperatures may have different molecular packing, thereby affecting the cross-linking efficiency of diacetylene moieties due to the topochemical nature of the polymerization. Large defects could be a consequence of the formation of domains, in which molecules cannot be polymerized efficiently and are thus removed by the subsequent SDS treatment. However, this possibility is unlikely due to the following experimental results. First, the residual film thickness after the SDS treatment was nearly identical for both samples. The film thickness measured by AFM was 4.8 ( 0.4 nm for samples prepared at 28 °C and 4.6 ( 0.5 nm for the samples prepared at 16 °C. The average thickness measured by ellipsometry (on silicon wafers) was identical for both conditions (4.4 ( 0.2 nm). Second, no significant differences in the UV absorption spectra from conjugated polymer backbones were observed among two samples (see the Supporting Information). Both samples showed spectral features of so-called “red films” having limited conjuga-

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tion length of the diacetylene backbone.19,20 This is attributed to the disorder in diacetylene backbones imposed by the packing constraints of diacylglycerol units in DiynePC molecules.15 The similarity of UV absorption spectra suggests that the molecular order in both films is similar. Another potential mechanism for the observed temperature effect is thermal polymerization of DiynePC. Although we are not aware of any systematic study on thermal polymerization of diacetylene phospholipids, we suppose that it should not play an important role in the film morphologies in the present samples from the fact that thermal polymerization is kinetically hindered for diacetylene derivatives having bulky side groups.32 Diacetylene phospholipids have a rather bulky glycerol backbone, and the packing of diacetylene units should be less favorable for thermal polymerization compared with single-chain diacetylene fatty acids. The fact that we did not observe any significant influence of thermal polymerization in the annealing experiments also supports the premise that thermal polymerization is not playing a major role in the polymer morphologies observed at two temperatures. Therefore, the most plausible mechanism for the observed defect formation is the destabilization of polymerized bilayers on the surface due to the constraints in film curvature and area. Formation of tubular structures from diacetylene bilayers was reported in previous literature, indicating that polymerized diacetylene membranes have a finite spontaneous curvature.33,34 Furthermore, previous studies reported that DiynePC expanded in the lateral direction upon polymerization, if a monolayer was polymerized on the water surface keeping the surface pressure constant.23 The mechanical stress within the adsorbed bilayer should cause partial detachment of the film from the surface. In fact, the elongated appearance of the defects described as “channel-like” may very well be the consequence of mechanical stress in the polymerized bilayer film. Similar failures of materials have been reported for bulk polydiacetylenes. It is plausible that the destabilization effects depend on the size of bilayer domains, with larger domains being more drastically affected by the curvature and area constraints upon polymerization. In summary, we have found that the homogeneity of polymerized bilayers from a diacetylene-containing phospholipid (DiynePC) was very sensitively influenced by the film deposition temperature and alternatively by the annealing/quenching

Figure 7. (a) Contact mode AFM height (z-scale 50 nm) and (b) friction difference image (friction forces [a.u.] increase from dark to bright contrast) of patterned bilayer deposited at 16 °C acquired in water at RT after SDS treatment.

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Figure 8. (a) Contact mode AFM force volume height (z-scale 50 nm) and (b) force volume image (pull-off forces [zero to 6 nN] increase from bright to dark contrast) of patterned bilayer deposited at 16 °C and acquired in water at RT after SDS treatment. The difference in pull-off force between the AFM tip and the patterned surface is also evident in the overlaid force-displacement curves (acquired at the locations marked with a cross in the force volume image) shown in the lower panel c; the elevated bilayer region exhibited a pull-off force of ∼2.1 nN, while the glass substrate (inside the square shaped depressions) showed repulsive interactions only.

protocols. Whereas DiynePC bilayers (monomer) deposited at a temperature ∼4 °C below the triple point temperature of DiynePC monolayers (∼20 °C) formed homogeneous polymeric bilayers, those deposited at a temperature ∼8 °C higher than the triple point temperature showed a markedly increased number of line defects. The differences were attributed to the domain structures in the monolayer on water surface. Since the progress of polymerization detected by the polymer backbone conjugation (UV/visible absorption spectra) and the residual film thickness (AFM and ellipsometry) were unaffected by the film deposition temperatures, the domain size, rather than the molecular packing in each domain, seems to be playing a critical role. Changes in the spontaneous curvature and film area are the plausible source of destabilization and detachment of the films upon polymerization. The results from annealing experiments supported this conclusion. The morphological and chemical mapping by AFM demonstrated that the bilayer deposition, polymerization and subsequent removal of monomeric lipid work well, if monomeric (32) Eckhardt, H.; Prusik, T.; Chance, R. R. Macromolecules 1983, 16, 732736. (33) Yager, P.; Schoen, P. E. Mol. Cryst. Liq. Cryst. 1984, 106, 371-381. (34) Schnur, J. M.; Ratna, B. R.; Selinger, J. V.; Singh, A.; Jyothi, G.; Easwaran, K. R. K. Science (Washington) 1994, 264, 945-947.

bilayers were assembled below the triple point temperature. At the same time, the present results highlight the importance of film fabrication conditions for ensuring the homogeneity of polymerized bilayers in the micrometer scale, which is prerequisite for generating microarrays of lipid membranes useful for biomedical applications. Acknowledgment. We thank Ms. Maki Koike for her assistance in the preparation of patterned substrates and vesicle suspensions. This work has been supported by Promotion Budget for Science and Technology (AIST Upbringing of Talent in Nanobiotechnology Course) from the Ministry of Education, Science, Culture and Sports (MEXT), Grant-in-Aid for Scientific research from Japan Society for the Promotion of Science, Sekisui Chemical Grant Program (grant awarded to K.M.), and the Council for Chemical Sciences of The Netherlands Organization for Scientific Research (CW-NWO) in the framework of the Vernieuwingsimpuls program (grant awarded to H.S.). Supporting Information Available: UV/visible absorption spectra of ploymerized DiynePC bilayers (Figure S1). This information is available free of charge via the Internet at http://pubs.acs.org. LA701346X