Using Bicellar Mixtures To Form Supported and Suspended Lipid

New Mexico, Albuquerque, New Mexico 87131, Center for High Technology ... containing arrays of parallel troughs (170 nm wide, 380 nm deep, and 300 nm ...
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Using Bicellar Mixtures To Form Supported and Suspended Lipid Bilayers on Silicon Chips Reema Zeineldin,† Julie A. Last,§ Andrea L. Slade,§ Linnea K. Ista,† Paul Bisong,† Michael J. O’Brien,† S. R. J. Brueck,‡ Darryl Y. Sasaki,*,§ and Gabriel P. Lopez*,† Center for Biomedical Engineering and Department of Chemical & Nuclear Engineering, UniVersity of New Mexico, Albuquerque, New Mexico 87131, Center for High Technology Materials, UniVersity of New Mexico, Albuquerque, New Mexico, 87106, and Biomolecular Interfaces & Systems, Sandia National Laboratories, Albuquerque, New Mexico, 87185 ReceiVed March 27, 2006. In Final Form: June 22, 2006 Bicellar mixtures, planar lipid bilayer assemblies comprising long- and short-chain phosphatidylcholine lipids in suspension, were used to form supported lipid bilayers on flat silicon substrate and on nanotextured silicon substrates containing arrays of parallel troughs (170 nm wide, 380 nm deep, and 300 nm apart). Confocal fluorescence and atomic force microscopies were used to characterize the resulting lipid bilayer. Formation of a continuous biphasic undulating lipid bilayer membrane, where the crests and troughs corresponded to supported and suspended lipid bilayer regions, is demonstrated. The use of interferometric lithography to fabricate nanotexured substrates provides an advantage over other nanotextured substrates such as nanoporous alumina by offering flexibility in designing different geometries for suspending lipid bilayers.

Introduction Supported lipid bilayers (SLBs) have played an important role in the characterization of biological membranes,1,2 providing a platform to study the physical and mechanical properties of the lipid membrane, the molecular dynamics of membrane components, and the behavior of membrane-associated proteins. In biological studies, SLBs have been used to investigate cellular communication and adhesion, receptor-ligand binding, and biosensing.1,3-5 The archetypal SLB structure is a ∼5 ( 1 nm thick phospholipid membrane residing upon an ∼1 nm water layer that rests on a solid support.6,7 Typically, SLBs are formed on flat substrates, such as mica, glass, or silica, but a number of examples of SLBs on structured solid substrates have recently been reported. Microfabricated surfaces with lithographically derived features have been designed to partition the supported membrane into discrete and separate regions,8 to pattern the distribution of SLBs over a structured surface,9 or to facilitate lipid bilayer coating of microchannels.10 Lipid bilayers supported on microstructured arrays have also been used in Brownian ratchet devices for separating charged phospholipids.11 In addition, nanoporous alumina has been used as a support for bilayers to * To whom correspondence should be addressed. Phone: (505) 2774939; e-mail: [email protected] (G.P.L.). Phone: (925) 294-2922; e-mail: [email protected] (D.Y.S.). † Center for Biomedical Engineering and Department of Chemical & Nuclear Engineering, University of New Mexico. ‡ Center for High Technology Materials, University of New Mexico. § Sandia National Laboratories. (1) Sackmann, E. Science 1996, 271, 43-48. (2) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149-157. (3) Groves, J. T.; Dustin, M. L. J. Immunol. Methods 2003, 278, 19-32. (4) Anrather, D.; Smetazko, M.; Saba, M.; Alguel, Y.; Schalkhammer, T. J. Nanosci. Nanotechnol. 2004, 4, 1-22. (5) Cooper, M. A. J. Mol. Recognit. 2004, 17, 286-315. (6) Bayerl, T. M.; Bloom, M. Biophys. J. 1990, 58, 357-362. (7) Koenig, B. W.; Krueger, S.; Orts, W. J.; Majkrazk, C. F.; Berk, N. F.; Silverton, J. V.; Gawrisch, K. Langmuir 1996, 12, 1343-1350. (8) Cremer, P. S.; Yang, T. J. Am. Chem. Soc. 1999, 121, 8131-8131. (9) Nissen, J.; Jacobs, K.; Ra¨dler, J. O. Phys. ReV. Lett. 2001, 86, 1904-1907. (10) Yang, T.; Jung, S.-Y.; Mao, H.; Cremer, P. S. Anal. Chem. 2001, 73, 165-169. (11) van Oudenaarden, A.; Boxer, S. G. Science 1999, 285, 1046-1048.

examine lipid diffusion rates using electrochemical techniques and as a means to enhance lipid membrane surface area.12 An important issue in the development of SLBs for biosensing and biomembrane modeling studies is the decoupling of the membrane from substrate influences. The substrate can inhibit membrane fluidity by interacting with the lipid components and can also impair protein mobility, and thus function. Past research has focused on avoiding these limitations by extending the distance between the lower leaflet of the SLB and the substrate through the use of long hydrophilic spacers,13 tethering from soft polymer cushions,14 and the creation of undulated bilayers in porous alumina.15 Most recently, lipid bilayers have been suspended over micromachined holes16 and nanoscale pores.17 For microscale pores, the bilayers are prepared in a way identical to that for forming black lipid membranes (BLMs) and are thus termed micro-BLMs.18 On nanoscale porous anodic alumina substrates, lipid vesicle fusion has generated bilayer-coated pores that display gigaohm resistance with excellent single ion channel sensitivity for biosensing applications.19 Suspended lipid bilayers offer unique opportunities for the study of cell membranes and lipid bilayer physics as well as for development of new biosensing devices. The micro- and nanoBLM structures described above offer many advantages for device applications, but have some limitations with regard to studying membrane structure and dynamics because of the confinement and mobility of lipids and proteins to within the isolated suspended areas.20 Furthermore, the geometry of the structure is not amenable to fluid exchange beneath the suspended bilayer. (12) Marchal, D.; Boireau, W.; Laval, J. M.; Moiroux, J.; Bourdillon, C. Biophys. J. 1998, 74, 1937-1948. (13) Lang, H.; Duschl, C.; Vogel, H. Langmuir 1994, 10, 197-210. (14) Sinner, E. K.; Wolfgang, K. Curr. Opin. Chem. Biol. 2001, 5, 507-711. (15) Gaede, H. C.; Luckett, K. M.; Polozov, I. V.; Gawrisch, K. Langmuir 2004, 20, 7711-7719. (16) Ogier, S. D.; Bushby, R. J.; Cheng, Y.; Evans, S. D.; Evand, S. W.; Jenkins, T. A.; Knowles, P. F.; Miles, R. E. Langmuir 2000, 16, 5696-5701. (17) Hennesthal, C.; Steinem, C. J. Am. Chem. Soc. 2000, 122, 8085-8086. (18) Ro¨mer, W.; Lam, Y. H.; Fischer, D.; Watts, A.; Fischer, W. B.; Go¨ring, P.; Wehrspohn, R. B.; Go¨sele, U.; Steinem, C. J. Am. Chem. Soc. 2004, 126, 16267-16274. (19) Drexler, J.; Steinem, C. J. Phys. Chem. B 2003, 107, 11245-11254. (20) Ro¨mer, W.; Steinem, C. Biophys. J. 2004, 86, 955-965.

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Herein, we examine the use of lipid bicellar mixtures, bilayer architectures comprising long- and short-chain phospholipids in aqueous suspension, which have been used to incorporate and study transmembrane proteins,22,23 to fabricate suspended bilayer membranes on a new type of nanotextured substrate. Bicellar mixtures, prepared by adding short-chain phosphatidylcholine (PC) lipids (six to eight carbon chain length) to long-chain PC lipids (carbon chain length g14), can form several morphological lipid assemblies in aqueous suspensions including planar lipid assemblies, such as bicelles, long ribbonlike micelles (or quasicylindrical micelles), and branched, flattened cylindrical micelles.24,25 The factors that determine the structural form of the bicellar mixture are temperature, the molar ratio of long-chain to short-chain PC (q ratio), and the lipid weight fraction in suspension (CL). Between the values of 0.5 e q e 4 and 1% e CL e 40%, as either the temperature or the q ratio increases, increasingly larger planar bilayer assemblies form in accordance with the morphological sequence mentioned above. On the other hand, at high temperatures or q ratios more complex nonplanar assemblies form, including perforated lamellar sheets and multilamellar vesicles.24,26,27 In general, suspensions containing the planar lipid assemblies formed by the bicellar mixtures possess great potential as model membrane mimics, because their planar lipid bilayer structure resembles that of biological membranes. Short-chain PCs are desirable detergents for solubilizing membrane proteins,28 and the formation of bicellar mixtures containing transmembrane proteins has been demonstrated to be a versatile tool in structural studies. In this paper we present a new architecture for suspended lipid bilayers formed from bicellar mixtures and prepared on nanotextured oxidized silicon substrata. The nanotextured surface, which consists of a high-density array of nanoscale troughs, is formed by interferometric lithography.21 The goals of this study are (1) to evaluate the ability of planar lipid bilayer assemblies of bicellar mixtures in suspension to form SLBs on flat substrates, (2) to obtain insight into the organization of these assemblies on a solid support by confocal microscopy and atomic force microscopy (AFM) and evaluate the mixing behavior of these hydrophobically mismatched lipids in SLBs, and (3) to investigate formation of a suspended lipid bilayer using these planar assemblies on a nanotextured, oxidized silicon wafer. The formation of lipid bilayers on nanotextured substrates using bicellar mixtures containing membrane proteins will enable using the suspended lipid bilayers as electrophoretic or chromatographic media, thus facilitating separation and examination of membrane components. The nanofabrication method used here, based on interferometric lithography, is particularly wellsuited for the formation of “artificial gels”29 because it is amenable to inexpensive, rapid patterning of large numbers of nanostructures over macroscopic areas.21 (21) O’Brien, M. J.; Bisong, P.; Ista, L. K.; Rabinovich, E. M.; Garcia, A. L.; Sibbett, S. S.; Lopez, G. P.; Brueck, S. R. J. J. Vac. Sci. Technol., B 2003, 21, 2941-2945. (22) Sanders, C. R.; Oxenoid, K. Biochim. Biophys. Acta: Biomembr. 2000, 1508, 129-145. (23) Marcotte, I.; Auger, M. Concepts Magn. Reson. A 2005, 24A, 17-37. (24) van Dam, L.; Karlsson, G.; Edwards, K. Biochim. Biophys. Acta 2004, 1664, 241-256. (25) Harroun, T. A.; Koslowsky, M.; Nieh, M.-P.; de Lannoy, C.-F.; Raghunathan, V. A.; Katsaras, J. Langmuir 2005, 21, 5356-5361. (26) Nieh, M.-P.; Raghunathan, V. A.; Glinka, C. J.; Harroun, T. A.; Pabst, G.; Katsaras, J. Langmuir 2004, 20, 7893-7897. (27) Triba, M. N.; Warschawski, D. E.; Devaux, P. F. Biophys. J. 2005, 88, 1887-1901. (28) Hauser, H. Biochim. Biophys. Acta 2000, 1508, 164-181. (29) Turner, S. W.; Perez, A. M.; Lopez, A.; Craighead, H. G. J. Vac. Sci. Technol., B 1998, 16, 3835-3840.

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Experimental Section Materials. 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and 1,2-diheptanoyl-sn-glycero-3-phosphocholine (DHPC) were obtained from Avanti Polar Lipids, Inc. (Alabaster, AL). N-(6Tetramethyl-rhodamine-thiocarbamoyl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (TRITC-DHPE), and N-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3propionyl)-1,2-dihexanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (PE-BODIPY), were purchased from InvitrogenMolecular Probes (Carlsbad, CA). Silicon Wafers. Thermally oxidized flat silicon wafers were obtained from NOVA Electronic Materials, Ltd. (Carrollton, TX). Nanotextured silicon wafers were fabricated as previously described using interferometric lithography.21 All wafers were cleaned by a quick dip in piranha solution (1 part 30% H2O2, 2 parts H2SO4 by volume), followed by thorough rinsing with ultrapure water. Formation of Bicellar Mixtures. Solutions of long- and shortchain PC lipids were prepared separately using lipid solutions in chloroform that were dried by nitrogen gas followed by vacuum for 10 min. A multilamellar vesicle solution of long-chain PC in 75 mM phosphate buffer, pH 7.0 was prepared from a 72 mM DPPC containing 0.5 mol % TRITC-DHPE in chloroform. A micellar solution of short-chain PC in 75 mM phosphate buffer, pH 7.0 was prepared from a 104 mM solution of DHPC containing 0.1 mol % of PE-BODIPY in chloroform. Bicellar mixtures were then formed at room temperature by adding the DHPC solution to the DPPC solution while mixing to yield final concentrations of 10 and 28 mM, respectively (q ratio ) 2.8, and CL of 2.5 wt %). The mixture was hydrated by storing at 4 °C for 19-24 h before use. We chose to prepare the bicellar mixtures at a q ratio of 2.8, and at room temperature, to form planar bilayer assemblies. Dynamic light scattering measurements using a Microtrac-S3000 laser particle size analyzer (Microtrac, Inc., North Largo, FL) indicated that the bicellar assemblies consisted of two populations with average lengths of 750 ( 250 nm and 5 ( 3 µm. Literature reports on assemblies formed under similar conditions but using a mixture of 14-carbon PC and 6-carbon PC include one study measuring a length on the order of microns24 and another estimating a length of >300 nm.26 Formation of Lipid Bilayers Using Bicellar Mixtures. Lipid bilayers were formed on the silicon dioxide surface by placing a 40 µL drop of the lipid suspension in a glass Petri dish and then placing the wafer on the drop for 10 min. The Petri dish was then filled with ultrapure water and the assembly was rinsed by submerging in a larger crystallization dish filled with ultrapure water with gentle shaking; rinsing was repeated two more times. Imaging of Lipid Bilayers Using Confocal Microscopy. A wafer with formed lipid bilayer was inverted while submerged, and transferred into a 2 cm × 2 cm square container with ultrapure water in it. A coverslip was placed on top of the wafer, and excess water was removed. Confocal images were generated using a Zeiss LSM 510 confocal microscope (Carl Zeiss MicroImaging Inc., Thornwood, NY), with excitation at 488 nm using an argon laser and at 543 nm using a helium-neon laser. Imaging of Lipid Bilayers Using Atomic Force Microscopy (AFM). A Si sample was transferred to the AFM liquid cell by placing the O-ring of the liquid cell on the sample while it was under water. The O-ring was then clamped to the sample using tweezers and carefully removed from the water so that the sample surface remained submerged under water. The bottom of the sample was carefully dried with a towel, and the sample was attached to an AFM puck on the scanner with double-sided tape. The liquid cell was then quickly assembled and filled with deionized water. AFM imaging of the SLBs was performed with a Nanoscope IIIa Multimode microscope (Digital Instruments, Santa Barbara, CA), with images acquired in solution with tapping mode using a commercially available liquid cell (Digital Instruments). Images were collected with the E-scanner operating at a scan rate of 2 Hz using a 120 µm oxide sharpened silicon nitride V-shaped cantilever having a nominal spring constant of 0.35 N/m.

Suspended Lipid Bilayers on Silicon Chips

Figure 1. SLB on a flat silicon substrate prepared with a 2.8:1 DPPC (with 0.1 mol % BODIPY-PE)/DHPC (with 0.5% TRITCDHPE) bicellar mixture. (A) Confocal microscopy images after lipid bilayer formation. The left inset is of Rhodamine fluorescence, the right inset is of BODIPY fluorescence, and the main image is a superimposition of the inset images. (B) Tapping mode AFM image after lipid bilayer formation. The black arrow points to a region rich in DPPC, and the green arrow points to a region rich in DHPC. The section analysis (bottom) shows the topographic profile along the dashed line, revealing a bilayer height of 59 Å and a DHPC-DPPC height difference of 14 Å. Scale bar ) 1 µm.

Results and Discussion Examination of Bicellar Mixtures-SLBs on Flat Substrate. To evaluate the structure and surface coverage of SLBs formed from the bicellar assemblies, an analysis was first performed on a flat (untextured) silicon wafer. Examination of the submerged assembly under a confocal microscope revealed the formation of a lipid bilayer fully covering the flat substrate with fluorescence from both the labeled long- and short-chain lipids distributed all over the substrate (Figure 1A). This distribution of fluorescence was similar to that obtained with egg-PC vesicles fused to silicon wafers (data not shown). Images of the 2.8:1 DPPC/DHPC bilayers on flat silicon revealed a continuous lipid bilayer composed of two phases, which were readily observed due to differences in the heights

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of the molecules (Figure 1B). The black arrow in Figure 1B points to a region rich in DPPC, while the green arrow points to a region rich in DHPC. The height difference between the DPPC and DHPC areas was measured to be ∼14 Å, which corresponds to the difference between the chain lengths of DHPC (7-carbon chain) and DPPC (16-carbon chain). The phase separation was due to immiscibility of the two phases stemming from the preferential interactions between the acyl chains of similar length in the fluid (DHPC) vs the gel phase (DPPC). Such phenomena have been well characterized by AFM with mixtures of PCs of different hydrocarbon lengths.30 A defect detected in the DPPC portion of the SLB shows a depth of ∼59 Å, corresponding to a depth of ∼49 Å for the formed SLB with an ∼10 Å underlying water layer, which is comparable to values found in the literature.7 Examination of Bicellar Lipid Bilayers on Nanotextured Substrate. The nanochannel structure in silicon was fabricated as previously described using interferometric lithography.21 The cross-sectional dimensions of the channels, as determined by scaning electron microscopy (SEM), indicate a tapered structure with a width of ∼175 nm at the top of the channels and ∼100 nm at the bottom, and with a depth of ∼380 nm (Figure 2A). The width of the ridges was ∼300 nm. Tapping mode AFM of the structure confirms the channel width of ∼175 nm (Figure 2B); however, the size of the imaging tip (50 nm radius) prevents an accurate measurement of the channel depth and tapered structure. A maximum depth of ∼100 nm was attained by AFM. Preparation of the SLB on the nanotextured silicon was done in a manner identical to that on the flat silicon. Confocal fluorescence microscopy of the surface with an adsorbed bilayer (Figure 3A) indicated that, similar to on flat substrate, the longand short-chain lipids were distributed all over the nanotextured surface. At higher resolution confocal fluorescence microscopy (Figure 3B) we see clear evidence of segregated distribution of the green and red fluorescence. This further provides support for our assumption that the green probe segregates with the short lipids and the red probe segregates with the long lipids. AFM imaging also indicated that the nanotextured silicon was coated with the lipid bilayer (Figure 3C), covering the tops of the ridges as a single bilayer and suspending across the channels. The bilayer topology was found to undulate with the same periodicity as the bilayer-absent wafer, but with a more shallow trough of ∼43 nm within the channel regions. Bilayer structural features were difficult to discern from the areas on the ridge tops due to the surface’s inherent roughness (see Figure 2B). However, the suspended regions provided areas removed from surface effects, permitting detailed examination of the bilayer. For example, within the dotted-line box of Figure 4, the dark blue arrow points to an area that is consistent with a feature rich in DPPC, while the pink arrow points to a region rich in DHPC based on height and surface area coverage. A hole in the lipid bilayer within the channel was also detected by AFM (light blue arrow in center of Figure 4), suggesting that the suspended portion of the lipid bilayer is not immediately supported by lipid material trapped within the channels of the silicon wafer; further work is needed to confirm this. While a height measurement cannot be taken from the bilayer suspended above the channel, the presence of a single bilayer on the flat silicon chip (Figure 1B) indicates that a single bilayer covers the channels. In addition, partial multilayers were never observed on either the flat or corrugated silicon substrates. These results demonstrate the (30) Tokumasu, F.; Jin, A. J.; Feigenson, G. W.; Dvorak, J. A. Biophys. J. 2003, 84, 2609-2618.

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Figure 2. Imaging of a nanotextured silicon wafer typical of wafers used in this study. (A) SEM image of nanotextured silicon after etching and oxidation reveals a channel depth of 380 nm, a channel width of 175 nm, and a ridge width of 300 nm. (B) Tapping mode AFM image (scale bar ) 375 nm) and section analysis (bottom) of the nanotextured silicon show a topographic profile (along the dashed line) with a channel depth of ∼91 nm and width of ∼170 nm.

feasibility of using bicellar assemblies to form suspended single lipid bilayers on nanochannel architectures. Our findings contrast recent work where chemical modification of the substrate is required to facilitate suspended bilayer formation,16,17 or where a double lipid bilayer lobe is formed on the surface of a single lipid bilayer spreading along the bottom of microgrooves.31 Previous studies using lipid vesicles to form SLBs on flat supports found slow formation of SLBs when performed at a temperature close to or below the transition temperature (Tm) of single or mixed long-chain PC lipids.32,33 This was found to be a result of slower vesicle rupture, but not vesicle adsorption to the support. Since our study used planar lipid bicellar assemblies, instead of vesicles, no postprocessing of the bilayer structure is necessary. All that is required for these assemblies is adsorption to the support followed by fusion between (31) Suzuki, K.; Masuhara, H. Langmuir 2005, 21, 6487-6494. (32) Seantier, B.; Breffa, C.; Fe´lix, O.; Decher, G. Nano Lett. 2004, 4, 5-10. (33) Beckmann, M.; Nollert, P.; Kolb, H.-A. J. Membr. Biol. 1998, 161, 227233.

Figure 3. SLB on a nanotextured silicon substrate prepared with a 2.8:1 DPPC (with 0.1 mol % BODIPY-PE)/DHPC (with 0.5% TRITC-DHPE) bicellar mixture. (A) Confocal microscopy images after lipid bilayer formation, and with higher resolution in (B). The right inset is of Rhodamine fluorescence, the left inset is of BODIPY fluorescence, and the main image is a superimposition of the inset images. (C) Tapping mode AFM of a nanotextured silicon substrate after lipid bilayer formation (scale bar ) 375 nm). The section analysis (bottom) shows the topographic profile along the dashed line, revealing a depth in the center of the channel of 43 nm.

adsorbed bilayer patches to form a continuous SLB. By eliminating the slow step of vesicle rupturing at low temperatures

Suspended Lipid Bilayers on Silicon Chips

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Figure 4. Tapping mode AFM image of a lipid bilayer on a nanotextured silicon substrate. The white box highlights an area of the bilayer over the channel where both areas rich in DPPC and areas rich in DHPC are evident. Within the box, the dark blue arrow points to a region rich in DPPC and the pink arrow points to a region rich in DHPC. The light blue arrow points to a hole in the lipid bilayer. Scan size ) 750 nm. Scale bar ) 175 nm.

for vesicle fusion, using planar bicellar assemblies may reduce the dependence of SLB formation on the temperature of the surrounding environment. It has been reported that discoidal phospholipid nanoparticles, which comprise a lipid bilayer, adsorb to mica and remain intact as a monolayer of disks on mica.34 However, upon application of high force loads, these disks fuse to form lipid bilayer domains on mica.34 In our study, the large diameter of the planar bicellar assemblies probably facilitated the SLB formation without application of any force. Furthermore, our use of planar bicellar assemblies with diameters larger than the channel width facilitated the suspension of a lipid bilayer on the nanotextured support. Figure 5 shows three-dimensional representations of the AFM images of the uncoated and bilayer-coated nanochannel structures. From the images it is evident that a relatively smooth and continuous bilayer was supported and suspended across the structure. The undulations have radii of curvature of ∼175 nm in both the crests and troughs of the supported and suspended areas, respectively. Sagging of the suspended bilayers has been observed previously with bilayers draped over nanoscale pores.35 It is not likely that this bilayer structure is an artifact of the imaging process, but rather it is a product of the interaction of the bilayer with the tops of the channel walls. Our ability to consistently and reproducibly attain the same images of the suspended bilayer structures with a sagging depth of ∼40 nm regardless of different tips used, and the fact that the imaging is clear and resolved in this region, attests to a stable membrane structure. Strong coupling of the bilayer to the silicon dioxide surface and the rounded edges of the tops of the channel walls may cause the bilayer to come off the tops of the ridges at a downward pitch, producing the undulated bilayer structure. That conformal coating of the bilayer was not observed may be a product of the width of the channel structure relative to the size of the bicellar assemblies and the preferred curvature of the lipid membranes, which would be rather acute at the corner of the tops of the channel walls. (34) Bayburt, T. H.; Grinkova, Y. V.; Sligar, G. Nano Lett. 2002, 2, 853-856. (35) Hennesthal, C.; Drexler, J.; Steinem, C. ChemPhysChem 2002, 10, 885889.

Figure 5. Three-dimensional representations of AFM images of a nanotextured silicon substrate. (A) Three-dimensional representation of the substrate before application of the lipid mixture. (B) Threedimensional representation of the substrate after application of the 2.8:1 DPPC:DHPC bicellar mixture. Scan size ) 1.5 µm.

Conclusion This was the first demonstration of SLB formation using bicellar mixtures. Furthermore, preparation of suspended lipid bilayers on a nanotextured silicon substrate using these lipid bicellar mixtures was successfully demonstrated. The topography of the bilayer undulated in registry with the corrugation of the underlying silicon substrate. However, the suspended bilayer areas were sufficiently stable to allow for nanoscale imaging of bilayer structural features, demonstrating feasibility of probing nanoscale structures in suspended bilayer architectures. This study presents the use of interferometric lithography to fabricate nanotexured substrates for suspending lipid bilayers. Interferometric lithography has the advantage of offering flexibility in design of different geometries for suspending lipid bilayers when compared, for example, to nanoporous alumina. We are introducing a preliminary step to the formation of biomimetic SLBs in which the nanotextured silicon substratum provides a geometry for rapid fluid exchange under the suspended portion of the bilayer upon introducing or withdrawing a fluid at either end of the troughs. Another advantage of the nanotextured silicon substratum is that it offers an open membrane architecture where the mobility of lipids is allowed within the suspended lipid bilayer over the troughs, thus permitting free access to multiple membrane components for dynamic assembly of ion channels or membrane receptors. The ability of short-chain lipids to solubilize membrane proteins is ideal for incorporating membrane proteins into bicellar mixtures, thus facilitating the examination of proteins by the subsequent formation of an SLB. Although there are many advantageous properties associated with bicellar mixtures for membrane protein sequestration and

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characterization, to our knowledge there are no previous reports of the use of such systems to form SLBs. The bilayer-substrate architecture presented may serve as an ideal model membrane to mimic the cytoplasmic compartment, allowing for the study of membrane proteins with nanoprobes above and below the membrane. The expected increased fluidity of the lower leaflet of the lipid bilayer in the suspended regions, as well as the lack of steric hindrance of the intercellular protein domains, will result in improved mobility of incorporated proteins, and will eliminate undesired interactions that inactivate some transmembrane proteins. This system employing bicellar mixtures may have great potential for use in proteomics and in studying cellular interactions with membrane proteins and intracellular signaling molecules. Acknowledgment. This research was supported in part by the National Science Foundation (SENSORS: CTS 0332315,

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NIRT: EEC 0210835, and NIRT: CTS 0404124), the Army Research Office (DAAD19-03-1-0173), and the Division of Materials Science and Engineering in the Department of Energy Office of Basic Energy Sciences through the Center for Integrated Nanotechnologies (CINT). Sandia is a multiprogram laboratory operated by Sandia Corporation, a Lockheed Martin Company, for the United States Department of Energy’s National Nuclear Security Administration under Contract No. DE-AC04-94AL85000. Confocal images in this paper were generated in the UNM Cancer Center Fluorescence Microscopy Facility, which received support from NCRR 1 S10 RR14668, NSF MCB9982161, NCRR P20 RR11830, NCI R24 CA88339, NCRR S10 RR19287, NCRR S10 RR016918, the University of New Mexico Health Sciences Center, and the University of New Mexico Cancer Center. LA060817R