Nonequilibrium Patterns of Cholesterol-Rich Chemical

May 4, 2006 - Department of Applied Science, UniVersity of California, DaVis, California 95616, ... Chemistry, UniVersity of California, and Lawrence ...
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Langmuir 2006, 22, 5374-5384

Nonequilibrium Patterns of Cholesterol-Rich Chemical Heterogenieties within Single Fluid Supported Phospholipid Bilayer Membranes Annapoorna R. Sapuri-Butti, Qijuan Li, Jay T. Groves, and Atul N. Parikh* Department of Applied Science, UniVersity of California, DaVis, California 95616, and Department of Chemistry, UniVersity of California, and Lawrence Berkeley National Laboratory, Berkeley, California 94720 ReceiVed August 18, 2005. In Final Form: January 7, 2006 We have developed a simple method to introduce cholesterol- and sphingomyelin-rich chemical heterogeneities into controlled densities and concentrations within predetermined regions of another distinct fluid phospholipid bilayer supported on a solid substrate. A contiguous primary phasesa fluid POPC bilayer displaying a well-defined array of lipid-free voids (e.g., 20-100 µm squares)swas first prepared on a clean glass surface by microcontact printing under water using a poly(dimethylsiloxane) stamp. The aqueous-phase primary bilayer pattern was subsequently incubated with secondary-phase small unilamellar vesicles composed of independent chemical compositions. Backfilling by comparable vesicles resulted in gradual mixing between the primary- and secondary-phase lipids, effacing the pattern. When the secondary vesicles consisted of phase-separating mixtures of cholesterol, sphingomyelin, and a phospholipid (2:1:1 POPC/sphingomyelin/cholesterol or 1:1:1 DOPC/sphingomyelin/cholesterol), well-defined spatial patterns of fluorescence, chemical compositions, and fluidities emerged. We conjecture that these patterns form because of the differences in the equilibration rates of the secondary liquid-ordered and liquid-disordered phases with the primary fluid POPC phase. The pattern stability depended strongly on the ambient-phase temperature, cholesterol concentration, and miscibility contrast between the two phases. When cholesterol concentration in the secondary vesicles was below 20 mol %, secondary intercalants gradually diffused within the primary POPC bilayer phase, ultimately dissolving the pattern in several minutes and presumably forming a new quasi-equilibrated lipid mixture. These phase domain micropatterns retain some properties of biological rafts including detergent resistance and phase mixing induced by selective cholesterol extraction. These patterns enable direct comparisons of cholesterol- and sphingomyelin-rich phase domains and fluid phospholipid phases for their functional preferences and may be useful for developing simple, parallelized assays for phase and chemical composition-dependent membrane functionalities.

Introduction A growing body of evidence now suggests that biological membranes manage their vast chemical diversity by sorting into specialized compartments or microdomains.1-3 A pervasive class of membrane microdomains that exhibit a significant enrichment of cholesterol and sphingolipids in their material compositions, which are broadly referred to as lipid rafts, has received considerable attention in recent years.4,5 Interest in these microstructures stems from their possible role in a very broad range of important biological processes. These now include a vast class of signaling pathways, cell adhesion and migration, synaptic transmission, cytoskeletol organization, membrane transport, protein sorting, and apoptosis during normal cellular homeostasis.6,7 In addition, cholesterol- and sphingolipid-rich phase domains have also been suggested to serve as functional hot spots for bacteria, viruses, and toxins as well as provide a microenvironment for prion formation and amyloid aggregation.8 An early indication of the existence of membrane phase domains was provided by indirect evidence based on observations that specialized lipid fractions composed predominantly of sphingomyelin, glycosphingolipids, cholesterol, selected membrane proteins, and most likely saturated phospholipids were * Corresponding author. E-mail: [email protected]. (1) Jacobson, K.; Sheets, E. D.; Simson, R. Science 1995, 268, 1441-1442. (2) Vaz, W. L. C.; Almeida, P. F. F. Curr. Opin. Struct. Biol. 1993, 3, 482488. (3) Glaser, M. Curr. Opin. Struct. Biol. 1993, 3, 475-481. (4) Simons, K.; Ikonen, E. Science 2000, 290, 1721-1726. (5) Simons, K.; Ikonen, E. Nature 1997, 387, 569-572. (6) Simons, K.; Toomre, D. Nat. ReV. Mol. Cell Biol. 2000, 1, 31-39. (7) Simons, K.; Ehehalt, R. J. Clin. InVest. 2002, 110, 597-603. (8) Brown, D. A.; London, E. Annu. ReV. Cell DeV. Biol. 1998, 14, 111-136.

highly resistant to extraction using cold nonionic detergents.9,10 These observations led to the hypothesis that detergent-resistant microdomains (DRMs) arise primarily from the distinct microdomains on the plasma membranes that were enriched in DRM constituents. More recently, novel applications of Forster resonance energy transfer (FRET) and multiphoton microscopy methods have revealed the presence of microdomains within live cells.11-13 A central tenet of the raft model is the existence of liquid-ordered (lo) supramolecular dynamic assemblies enriched in sphingolipids and cholesterol surrounded by a phospholipidrich liquid-disordered (ld) membrane phase.1,4 It also proposes that raft domains preferentially incorporate selected classes of proteins, including glycosylphosphatidylinositol (GPI)-anchored proteins, doubly acylated peripheral membrane proteins, cholesterol-linked proteins, and some transmembrane proteins.1,4 It has been further suggested that the rafts form from the molecular packing requirements and equilibrium phase behavior of lipid mixtures.14-17 In particular, it is believed that the presence of extended acyl chains in sphingolipids, coupled with large polysaccharide headgroups, allows for the tight intercalation of cholesterol in their intermolecular free volume. As a result, the sphingolipid-cholesterol mixture phase separates from other unsaturated lipids in the membrane, presenting a notably denser local molecular environment. Thus, rafts form distinct liquid(9) Brown, D. A.; Rose, J. K. Cell 1992, 68, 533-544. (10) Shogomori, H.; Brown, D. A. Biol. Chem. 2003, 384, 1259-1263. (11) Gaus, K.; Gratton, E.; Kable, E. P. W.; Jones, A. S.; Gelissen, I.; Kritharides, L.; Jessup, W. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 15554-15559. (12) Varma, R.; Mayor, S. Nature 1998, 394, 798-801. (13) Kenworthy, A. K.; Edidin, M. J. Cell Biol. 1998, 142, 69-84.

10.1021/la052248d CCC: $33.50 © 2006 American Chemical Society Published on Web 05/04/2006

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ordered (lo) regions, dispersed within the liquid-disordered (ld) surroundings of other unsaturated lipids. To validate these notions, model phospholipid membranes, including monolayers, giant unilamellar vesicles (GUVs), and supported lipid bilayers, are proving to be very useful.15,18,19 They offer flexibility and control of bilayer compositions and are amenable to quantitative characterization. Applications of these model systems are aimed at generating a physical-chemical basis that may be used to understand the fundamental structural and dynamical characteristics (sizes, lifetimes, and chemical compositions). In particular, by using model systems it is now well established that cholesterol interacts more strongly with SM than with unsaturated PC.20,21 It is also well understood that cholesterol has a condensing effect on the SM, resulting in a reduced effective molecular area. As a result, ternary mixtures comprising PC, SM, and chol are expected to exhibit liquidliquid phase separation forming chol- and SM-rich liquid-ordered (lo) phases distinct from liquid-disordered (ld) phase of predominatly unsaturated PC. Indeed, significant evidence is now available that suggests that lipid mixtures comprising chol, SM, and PC mixtures phase separate into coexisting lo and ld phases for a broad range of molecular compositions15,22 In this regard, recent studies have established that these systems exhibit significant disparities that are strongly dependent on the configuration of the model system used. For instance, the morphologies and sizes of lo and ld coexisting phases, lo phase mobilities, and their relative equilibration rates and dynamics appear to be strongly influenced by the type of model system employed.15,23 One of the most controversial issues in this regard relates to the sizes of the lo phase domains.15 For example, GUVs, Langmuir monolayers, and asymmetric composition bilayers have all been used to determine the sizes and phase topologies of coexisting lo and ld phases. Many studies using GUVs report the formation of mobile circular or oval-shaped lo phase domains that are several micrometers (2-10 µm) in linear dimensions.24-27 By contrast, supported membranes formed by vesicle deposition procedures have been shown in many reports to yield submicroscopic, nanometer-scale domain morphologies using scanning probe microscopy techniques28-33 that are not resolved by (14) Radhakrishnan, A.; Anderson, T. G.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 12422-12427. (15) Simons, K.; Vaz, W. L. C. Annu. ReV. Biophys. Biomol. Struct. 2004, 33, 269-295. (16) Brown, D. Int. J. Med. Microbiol. 2002, 291, 433-437. (17) Brown, D. A.; London, E. J. Membr. Biol. 1998, 164, 103-114. (18) Mukherjee, S.; Maxfield, F. R. Annu. ReV. Cell DeV. Biol. 2004, 20, 839-866. (19) London, E. Curr. Opin. Struct. Biol. 2002, 12, 480-486. (20) Ohvo-Rekila, H.; Ramstedt, B.; Leppimaki, P.; Slotte, J. P. Prog. Lipid Res. 2002, 41, 66-97. (21) Mattjus, P.; Slotte, J. P. Chem. Phys. Lipids 1996, 81, 69-80. (22) de Almeida, R. F. M.; Loura, L. M. S.; Fedorov, A.; Prieto, M. J. Mol. Biol. 2005, 346, 1109-1120. (23) Binder, W. H.; Barragan, V.; Menger, F. M. Angew. Chem., Int. Ed. 2003, 42, 5802-5827. (24) Korlach, J.; Schwille, P.; Webb, W. W.; Feigenson, G. W. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 8461-8466. (25) Stottrup, B. L.; Veatch, S. L.; Keller, S. L. Biophys. J. 2004, 86, 29422950. (26) Baumgart, T.; Hess, S. T.; Webb, W. W. Nature 2003, 425, 821-824. (27) Kahya, N.; Scherfeld, D.; Schwille, P. Chem. Phys. Lipid 2005, 135, 169-180. (28) Milhiet, P. E.; Vie, V.; Giocondi, M. C.; Le Grimellec, C. Single Mol. 2001, 2, 109-112. (29) Giocondi, M. C.; Vie, V.; Lesniewska, E.; Milhiet, P. E.; Zinke-Allmang, M.; Le Grimellec, C. Langmuir 2001, 17, 1653-1659. (30) Yuan, C. B.; Furlong, J.; Burgos, P.; Johnston, L. J. Biophys. J. 2002, 82, 2526-2535. (31) Rinia, H. A.; de Kruijff, B. FEBS Lett. 2001, 504, 194-199. (32) Rinia, H. A.; Snel, M. M. E.; van der Eerden, J.; de Kruijff, B. FEBS Lett. 2001, 501, 92-96. (33) Nicolini, C.; Thiyagarajan, P.; Winter, R. Phys. Chem. Chem. Phys. 2004, 6, 5531-5534.

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conventional fluorescence microscopy measurements. Moreover, they appear to exhibit very different miscibility characteristics from those of GUVs. From an experimental point of view and also from the vantage point of applications, it appears important that the development of the phase behavior of lipids in supported membranes, which is distinct from that of GUVs and Langmuir monolayers, is needed. This is because supported membranes provide a highly convenient platform for many membrane-mimetic applications (e.g., biosensors).34,35 Furthermore, they offer considerable ease in the detailed characterization of membrane dynamics and functions using a variety of surface-science-based techniques.36,37 In this article, we describe the development of a supported membrane configuration that incorporates raft-enriched regions within predetermined microscopic areas of a fluid phospholipid bilayer. The juxtaposition of fluid and cholesterol-rich phase domains within single contiguous bilayers affords comparative studies of composition-dependent membrane properties and functions. Our approach simply involves the spatially patterned deposition of an unsaturated fluid phospholipid bilayer using microcontact printing followed by the incorporation of phase-separating molecular mixtures from their vesicular precursors. We show that depending on cholesterol concentration, sample temperature, and degree of saturation of the phosphatidylcholine chain, the equilibration of cholesterol-rich phase domains can be kinetically arrested, forming long-lived geometric patterns of compositional heterogeneity within the contiguous fluid bilayer environment. These patterns of compositional domains appear to preserve some biochemical characteristics of cellular phase domains including cholesterol-extraction-induced dissolution and detergent insolubility. Experimental Section Materials. 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), 1,2-dioleoyl-sn- glycero-3-phosphocholine(DOPC), and ovine monosialoganglioside (brain, ovine-ammonium salt) (GM1) were purchased from Avanti Polar Lipids (Alabaster, AL). Sphingomyelin, cholesterol, fluorescein isothiocyanate-derivatized B subunits of cholera toxin, FITC-CTB, and methyl-β-cyclodextrin were purchased from Sigma (St. Louis, MO). The fluorescent probelabeled lipids, 1-oleoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3-phosphoethanolamine (NBD-PE), Texas red-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (Texas red-DHPE), and marina blue 1,2-dihexadecanoyl-sn-glycero3-phosphoethanolamine (marina blue DHPE) were from Molecular Probes (Eugene, OR). Phosphate buffer saline (PBS, pH 7.2, 154 mM NaCl, 1.54 mM KH2PO4, and 2.71 mM Na2HPO4) was from Gibco-Invitrogen Corporation (Grand Island, NY). Solvents and chemical reagents (chloroform, Triton X-100 detergent, and sulfuric acid) were from EM Science (Gibbstown, NJ), but hydrogen peroxide was from Fisher Chemicals (Fairlown, NJ). All organic solvents were HPLC grade. All chemicals were used without further purification. Organic-free deionized water of high resistivity (∼18.2 mΩ‚cm) was obtained by processing water through a Milli-Q Plus water system (model ZD40-11595, Bedford, MA) consisting of a reverse-osmosis deionization cartridge and an ion exchange/carbon purification system. Corning glass coverslips (no. 11/2, 22 mm2, Fisher HealthCare, Houston, TX) were used as substrates unless noted otherwise. Poly(dimethylsiloxane) (PDMS) stamps for microcontact printing were prepared using a Sylgard 182 elastomer kit from Dow Corning Corporation (Midland, MI). All silicon substrates used as masters for microcontact printing were (100)-oriented polished silicon wafers from Wafernet (San Jose, CA). (34) Cornell, B. A.; BraachMaksvytis, V. L. B.; King, L. G.; Osman, P. D. J.; Raguse, B.; Wieczorek, L.; Pace, R. J. Nature 1997, 387, 580-583. (35) Bayley, H.; Cremer, P. S. Nature 2001, 413, 226-230. (36) Sackmann, E. Science 1996, 271, 43-48. (37) Boxer, S. G. Curr. Opin. Chem. Biol. 2000, 4, 704-709.

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Preparation of Small Unilamellar Vesicles. Small unilamellar vesicles (SUVs) were prepared following a previously published procedure.38 Briefly, desired lipids,which were in chloroform, were first mixed together in predetermined mole ratios in a glass vial. The solvent was subsequently evaporated using a stream of nitrogen until dry, and the vial was kept under house vacuum for approximately 45 min to remove residual solvent that leaves behind a film of lipid at the bottom of the vial. The lipid film is then hydrated with filtered DI water (18.2 MΩ) and kept overnight at 4 °C. The total lipid concentration was 2 mg/mL, and the lipid solutions were used within 2 weeks. The hydrated lipids were sonicated for 4 min and were extruded with a miniextruder (Avanti Polar Lipids, Alabaster, AL) with 0.1 µm polycarbonate membrane filters (Whatman, Inc., Newton, MA); both were done at a temperature at least 10 °C above the transition temperature of the lipid or the lipid mixtures used. The lipid solution was passed through the extruder at least 19 times. SUVs were stored at 4 °C and used within 2 days. Typical experiments utilized POPC doped with 1 mol % Texas red-DHPE as the primary fluid phase. A typical secondary intercalating phase was a cholesterol-rich lipid mixture composed of constant 28 mol % sphingomyelin and varying ratios of cholesterol and POPC or 2:1:1 POPC/cholesterol/sphingomyelin or 1:1:1 DOPC/ cholesterol/sphingomyelin with 2% GM1 and 3% NBD-PE or 3 mol % marina blue-DHPE incubated with the POPC pattern. In terms of the percent and number of moles, we used the following: 2:1:1, 50% POPC (6.99 × 10-7 moles), 22% cholesterol (3.077 × 10-7 moles), 23% sphingomyelin (3.22 × 10-7 moles), 2% GM1 (2.80 × 10-8 moles), and 3% NBD-PE (4.18 × 10-8 moles) and 1:1:1, 33% DOPC (4.75 × 10-7 moles), 31% cholesterol (4.44 × 10-7 moles), 31% sphingomyelin (4.46 × 10-7 moles), 2% GM1 (2.87 × 10-8 moles), and 3% NBD-PE (4.28 × 10-8 moles). Precursor SUVs comprising fluid POPC and 1 mol % Texas red-DHPE were prepared by extruding at room temperature. Secondary SUVs were extruded at 40 °C. Several other compositions for the cholesterolrich raftlike phase and different fluid lipids for the primary phase were also used as specified in the Results section below. Substrate Preparation. Substrates (Corning glass coverslips) were prepared by immersing in a freshly prepared 4:1 (v/v) mixture of sulfuric acid and hydrogen peroxide for a period of 4 to 5 min maintained at ∼100 °C (Caution: this mixture reacts Violently with organic materials and must be handled with extreme care.) The substrates were then withdrawn using Teflon tweezers, rinsed immediately about eight times with copious amounts of deionized water, and stored under water. Cleaned substrates were used within 1 day of pretreatment. MicroContact Printing of Single Supported Membranes. Spatially defined patterns of primary phospholipid bilayers were created using a soft lithography technique involving the use of PDMS stamps in conjunction with supported phospholipid bilayers as the ink material. The PDMS stamps were prepared by adapting the now well-established replica molding method.39 The process begins by first preparing silicon masters displaying the desired patterns in a photoresist coating using a standard photolithography processes. Briefly, 4 in. silicon wafers were first prebaked on hot plate for ∼10 min at 110 °C and were vapor primed with hexadimethyldisilazane (HMDS). The wafers were then coated with Shipley 1813 positive photoresist by spinning at 3000 rpm to obtain an approximately 1.6-µm-thick layer of photoresist. The photoresist-coated wafers were vacuum baked on hot plate for 10 min at 110 °C and cooled and then exposed to ultraviolet light (λ ) 385 and 408 nm) through photomasks displaying the desired pattern of chrome on glass (Photoscience, Inc., Torrance, CA). The exposed silicon wafers were developed in MF-319 for ∼1 min and then rinsed in DI water and dried with a N2 gun. The development step dissolved the exposed photoresist. The unexposed photoresist was further stabilized by curing the photoresist pattern on a hot plate for 10 min at 110 °C.

The process resulted in relief patterns defined by the masks. Silicon masters so prepared were further primed with HMDS vapors. The latter step renders the master surface hydrophobic, allowing the facile separation of PDMS stamps from the master. Next, we poured a 9:1 mixture of Sylgard 182 (Dow Corning, MI.) elastomer and its curing agent onto the silicon masters, followed by curing at 75 °C for 2 h. Cured PDMS stamps were then separated from the silicon masters by gently peeling them off. The surfaces of PDMS stamps so derived are hydrophobic. The stamp surfaces were then rendered hydrophilic by exposing the textured stamp surfaces to ozonegenerating short-wavelength (λ ) 184-257 nm) ultraviolet radiation for approximately 90 s. Typically, PDMS patterns displaying elevated grids and depressed square features were used. The smallest feature size used was 20 µm × 20 µm squares separated by 20 µm edgeto-edge feature separation. Other sizes included square patterns of 50, 100, and 250 µm linear dimensions. Microcontact printing of the lipid bilayer was carried out by adapting previously published procedures.40-42 The PDMS stamp was UV oxidized for approximately 90 s to render the PDMS surface hydrophilic. Within 15 min of UV-assisted surface oxidation, the PDMS was brought into contact with a solution of 1:1 lipid vesicles/ 1X phosphate buffer saline for approximately 2 min. Without exposing the stamp to air, the stamp was immersed in DI water, and the aqueous phase was exchanged to remove any excess vesicles. Keeping the PDMS under water at all times, the inked PDMS was then brought into contact with a freshly cleaned glass substrate using a light weight (6-13 g depending on pattern geometry) for 15-20 s in water. The stamp was then separated from the glass slide under water, and the slide was stored for further use. Patterns so formed were tested for fluidity by FRAP (fluorescence recovery after photobleaching); once they were fluid, they were used within 2 h of preparation. Patterned bilayers so formed are stable for several days. Secondary Intercalation in Patterned Bilayers. Microcontact printed phospholipid bilayer patterns were incubated with an SUV solution of a cholesterol-rich lipid mixture. In a typical experiment, 2:1:1 POPC/cholesterol/sphingomyelin or 1:1:1 DOPC/cholesterol/ sphingomyelin with 2% GM1 and 3% NBD-PE or 3 mol % marina blue-DHPE was incubated with the POPC pattern. To facilitate optical discrimination, the secondary SUVs were doped with 3 mol % NBDPE (green) or 3 mol % marina blue-DHPE. The primary POPC patterns were doped with Texas red-DHPE. A 2 mg/mL solution of secondary SUVs (100 µL) kept at 37 °C comprising chol/SM/PC was introduced into the PBS solution of the patterned bilayer for about 2 to 3 min. The final concentration of secondary SUVs comprising chol/SM/PC in solution is 0.016 mg/mL. The sample was then rapidly rinsed with copious amounts of DI water and examined using epifluorescence microscopy. Gm1-Cholera Toxin Binding Assay. To determine the distribution of Gm1 in patterned bilayer samples, a Gm1-CTB binding assay was performed. Commercially acquired FITC-labeled CTB (MW ) 60 000 Daltons) was diluted to 0.5 mg/mL in PBS (which contains 20% protein to give a protein concentration of 0.1 mg/mL). A 5 µL aliquot of the CTB solution was then introduced into the 3 mL aqueous PBS phase of the micropatterned bilayer. The final CTB concentration was ∼2.7 × 10-9 M. After approximately 15 min of incubation, the sample was rinsed several times with 1X PBS. To facilitate the characterization of FITC-CTB (green) in the epifluorescence characterization, the bilayer samples were doped with TRDHPE (red) for the primary POPC bilayer phase and marina blueDHPE (blue) for the raft composition microphase. Triton X-100 Treatment. Triton X-100 treatment was carried out at room temperature and 4 °C to examine the detergent solubility of raftlike microdomains engineered within the fluid POPC phase. The bilayer samples comprising Texas red-DHPE-doped POPC and NBD-PE-labeled raftlike phases were treated with 0.012 (v/v) %

(38) Mayer, L. D.; Hope, M. J.; Cullis, P. R. Biochim. Biophys. Acta 1986, 858, 161-168. (39) Whitesides, G. M.; Ostuni, E.; Takayama, S.; Jiang, X. Y.; Ingber, D. E. Annu. ReV. Biomed. Eng. 2001, 3, 335-373.

(40) Sapuri, A. R.; Baksh, M. M.; Groves, J. T. Langmuir 2003, 19, 16061610. (41) Hovis, J. S.; Boxer, S. G. Langmuir 2001, 17, 3400-3405. (42) Hovis, J. S.; Boxer, S. G. Langmuir 2000, 16, 894-897.

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Langmuir, Vol. 22, No. 12, 2006 5377 method.45,46 Typically, a circular region of the fluorescent bilayer sample, ∼30-100 µm in diameter, was illuminated at high power continuously at the excitation wavelength for the fluorophore through a Plan Fluor ELWD 60× (NA, 0.70) objective for ∼2 min. The exposure bleaches a dark spot on the bilayer caused by the photoexcitation of the fluorophore followed by an irreversible chemical transformation effected by its reaction with oxygen dissolved in the ambient buffer. After photobleaching, images of fluorescence recovery in the bleached area were recorded at 60 s intervals. The subsequent lateral motion of unperturbed fluorophore lipids from the unbleached background into the bleached spot (and vice versa) is recorded in the recovery profiles. It has been previously established that the precise shape of the recovery curve can be used to qualitatively characterize the nature of the fluorophore motion.45,46 Furthermore, for diffusionlike motions, the measurements of the time required for the fluorescence intensity to recover to halfway (t1/2) between its immediate postbleach value and its long-time assymptotic value was used to estimate the diffusion coefficient, D, which is a measure of the fluidity of the bilayer environment. Specifically, we followed a method developed by Yguerabide et al.47 that approximates the solution of the 2D lateral diffusion equation using a modified Bessel function. Here, the experimental fluorescence intensity versus time data are replotted as reduced intensity versus time. Here, the reduced intensity is given by

Figure 1. Schematic depiction of the backfilling of stamped membrane bilayer patterns. Triton X-100 at room temperature and with 0.033% Triton X-100 at 4 °C.32 The samples were then examined in real time using epifluorescence imaging. Cholesterol Depletion Method. The bilayer samples comprising Texas red-DHPE-doped POPC and NBD-PE-labeled raftlike phases of 10% cholesterol, 28% sphingomyelin, 57% POPC, 2% GM1, and 3% NBD-PE in 3 mL of 1X PBS were treated with 300 µL of 10 mM methyl-β-cyclodextrin (MβCD) to give a 1 mM total concentration of MβCD in solution.43 Similarly, the bilayer samples were treated with higher-concentration MβCD by adding 300 µL of 100 mM methyl-β-cyclodextrin (MβCD) to give a 10 mM total concentration of MβCD in solution.44 Epifluorescence Microscopy. A Nikon eclipse TE2000-S inverted fluorescence microscope (Technical Instruments, Burlingame, CA) equipped with an ORCA-ER (model LB10-232, Hamamatsu Corporation, Bridgewater, NJ) or Retiga-1300 CCD camera (Technical Instruments, Burlingame, CA) and a mercury lamp as the light source was used to visualize all fluorescent samples. Two filter wheels, one containing a set of excitation and the other emission filters, were mounted in front of the light source and the CCD camera, respectively. An extra triple-band emitter was installed in the dichroic mirror cube to aid in focusing through the eyepiece. Typically, images taken used either a Plan Fluor 10X (NA, 0.25), Plan Fluor ELWD 20X (NA, 0.45), Plan Fluor ELWD 40X (NA, 0.60), or Plan Fluor ELWD 60X (NA, 0.70) objective (Nikon, Japan). High-resolution images were obtained using 100× (NA, 1.4) oil-immersion objectives. Images were stored and processed using simple PCI software (Compix, Inc., Cranberry Township, PA) augmented with a quantitative dynamic intensity analysis module. Fluorescence images taken with the Texas red filter set were assigned the color red, and images taken with FITC and UV filters were assigned the colors green and blue, respectively. Excitation and emission maxima for the probes used were 583/601 nm for TR-DHPE, 463/536 for NBDDHPE, and 365/460 nm for MB-DHPE. To characterize membrane fluidity, a simple method to assess fluorophore mobility within the membrane media was employed. We used microscopy-based fluorescence photobleach recovery measurements by adapting the circular spot photobleaching (43) Leventis, R.; Silvius, J. R. Biophys. J. 2001, 81, 2257-2267. (44) Lawrence, J. C.; Saslowsky, D. E.; Edwardson, J. M.; Henderson, R. M. Biophys. J. 2003, 84, 1827-1832.

I(red) )

I(t) - I(∞) I(0) - I(∞)

where I(t), I(0), and I(∞) correspond to fluorescence intensity at time t upon photobleaching, prebleaching, and long-time asymptotic recovery values, respectively. The plot is then used to estimate t1/2, which is then used to calculate D ) 0.22ro2/t1/2, where ro refers to the initial size of the photobleached spot. Next, we plotted the reduced fluorescence intensity against normalized time () Dt/ro2) using the estimated D values to compare the shape of the experimental normalized recovery curve with the theoretical curve47 to qualitatively assess if the experimental mobilities are diffusionlike. Finally, a comparison between the prebleach intensity and the asymptotic fluorescence intensity was used to assess the presence of any immobile fractions. Note that the exposure time (∼2 min) required in objectivebased photobleaching introduces some bilayer fluidity-dependent inaccuracies in the measurements of exact diffusion constants for probe lipids.

Results and Discussion Microcontact Printing Followed by Backfilling Provides a Useful Means to Engineer Bilayer Compositions. Our general strategy for introducing chemical and compositional heterogeneities within a single monocomponent-supported phospholipid bilayer is schematically illustrated in Figure 1. First, spatially patterned deposition of fluid primary-phase lipid bilayers on solid surfaces is achieved using microcontact printing or stamping. Second, the lipid-free pattern voids are backfilled using a secondary phospholipid phase (or a phase mixture) via subsequent exposure to secondary vesicles. This results in spatially defined incorporation of the secondary lipids, which become contiguous with the extant primary-phase bilayer. In this way, dissimilar lipid phases sharing a diffusive boundary can be organized within a single bilayer. Depending on the relative “miscibility” of the two juxtaposed phases, the phases can mix. In this manner, the combination of patterning and backfilling can be used to engineer material heterogeneity within fluid membranes. (45) Axelrod, D.; Koppel, D. E.; Schlessinger, J.; Elson, E.; Webb, W. W. Biophys. J. 1976, 16, 1055-1069. (46) Koppel, D. E.; Axelrod, D.; Schlessinger, J.; Elson, E. L.; Webb, W. W. Biophys. J. 1976, 16, 1315-1329. (47) Yguerabide, J.; Schmidt, J. A.; Yguerabide, E. E. Biophys. J. 1982, 40, 69-75.

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Figure 2. Pattern erasure by backfilling the lipid patterns with comparable secondary vesicles. Epifluorescence images of (a) a 20 × 20 µm2 membrane pattern comprising 1% Texas red-DHPE and 99% POPC and (b-e) a time-lapse sequence of the backfilling by a comparable composition vesicles composed of 2% GM1 and 98% POPC.

The application of microcontact printing for patterning fluid bilayers has previously been shown to produce discrete patches of phospholipid bilayers as well as contiguous bilayer phases possessing well-characterized voids within the interior.40-42 Feature sizes ranging from 1 to several hundred micrometers can be routinely achieved using this technique. We used PDMS stamps for the patterned deposition of a fluid bilayer comprising 20, 50, and 100 µm2 grid features (where lipid deposition occurred) with corresponding sizes of lipid-free square voids. An example of an epifluorescence emission image for fluid POPC bilayers doped with 1 mol % Texas red-DHPE is shown in Figure 2a. The high-contrast fluorescent pattern in Figure 2a shows dark features, essentially devoid of fluorescence emission, separated by a bright, homogeneous fluorescent background. A comparison of the bright-field image of the stamp and the corresponding epifluorescence image of the patterned bilayer reveals that the sizes and shapes of the fluorescent patterns are comparable replicas of the mask pattern. Furthermore, a simple adaptation of fluorescence recovery after photobleaching (FRAP) confirmed the fluidity of stamped bilayers to be comparable to that obtained by vesicle spreading methods (data not shown). Using microcontact printing, membrane patterns of various sizes, shapes, types of lipid, and fluorophores (used for visualization) can be reproducibly obtained. These observations are in conformity with earlier reports in the literature40-42 and indicate the formation of fluid-patterned phospholipid bilayers using microcontact printing. To manipulate bilayer compositions, we explored the possibility of backfilling the pattern voids by the subsequent exposure of patterned bilayers to a secondary vesicular solution.48,49 We first exposed the patterned POPC bilayers to SUVs of comparable compositions. The patterns were composed of a stamped POPC bilayer with square voids, which were 20 µm × 20 µm and separated by a 40 µm center-to-center distance. To allow discrimination between the secondary backfilling phase and the primary patterned bilayer, the incoming vesicles were unlabeled. The time lapse images shown in Figure 2 reveal that, upon (48) Yee, C. K.; Amweg, M. L.; Parikh, A. N. J. Am. Chem. Soc. 2004, 126, 13962-13972. (49) Yee, C. K.; Amweg, M. L.; Parikh, A. N. AdV. Mater. 2004, 16, 1184-+.

incubation, the initial nonfluorescent voids begin to acquire fluorescence from the surrounding primary phase and are quickly erased, leading to the homogenization of fluorophore intensity across the entire POPC surface, eliminating the pattern. These observations establish that the pattern voids can be accessed for secondary vesicle fusion, which subsequently erases the pattern by developing a contiguity with the primary bilayer phase. Spatial Patterning of Cholesterol-Rich Chemical Heterogeneities Within Fluid-Supported Bilayers. The ability to create predetermined patterns of voids within phospholipid bilayers, in conjunction with the ability to backfill, makes it possible to manipulate membrane compositions in a controlled manner. To explore this possibility, we introduced SUVs comprising mixtures of phospholipid (PC), sphingomyelin (SM), and cholesterol (chol) within the patterns of fluid PC bilayers. The data shown in Figure 3 show fluorescence images of mixed bilayers obtained when TR-DHPE-labeled fluid POPC patterns were backfilled with lipid mixtures comprising approximately a 2:1:1 mixture of POPC (55%), cholesterol (25%), and sphingomyelin (25%) doped with 3 mol % NBD-DHPE and 2% Gm1. Previous studies have established that vesicles of these and similar compositions when ruptured and spread on hydrophilic surfaces give rise to nanoscale phase separation in supported membranes, forming a cholesteroland SM-rich lo phase coexisting with a PC-rich ld phase.28-33 A typical epifluorescence image in Figure 3 reveals a sharp, checkerboard emission pattern consisting of “green” squares due to NBD-PE emission embedded within the “red” background associated with the TR-DHPE emission. The coloration is false and is used only to differentiate the emission in the red of the Texas red fluorophore from that in the green by NBD. The NBD emission pattern mirrors the initial pattern voids before backfilling was carried out. Several features of these data are noteworthy. First, the appearance of the fluorescence signal associated with the NBDPE (green) exclusively in the pattern-void regions confirms the accessibility of the voids for subsequent vesicle adsorption. The pattern-erasure experiments (Figure 2) established the ability to introduce lipids identical to the ones patterned that subsequently become contiguous with the existing membrane. The results in

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Figure 3. Engineered raftlike membrane heterogeneities. Typical (a) red channel, (b) green channel, and (c) composite red and green channel epifluorescence emission images for bilayers obtained by backfilling NBD-labeled raft-forming small unilamellar vesicles comprising 3% NBD-PE, 45% POPC, 25% cholesterol, 25% sphingomyelin, and 2% GM1 within a 20 × 20 µm2 stamped TR-DHPE-labeled POPC pattern.

Figure 4. Recognition of raft-partitioning GM1 by cholera toxin B5 subunits. Epifluorescence images of 20 × 20 µm2 stamped bilayer patterns (a) comprising 1% Texas red-DHPE and 99% POPC, (b) backfilled using raft-forming SUVs composed of 3% marina blue-DHPE, 45% POPC, 25% cholesterol, 25% sphingomyelin, and 2% GM1, and (c) upon incubation with FITC-labeled cholera toxin.

Figure 3 extend these observations by revealing that vesicles of markedly different compositions can also be used to backfill the patterns of voids. Second, the fluorescence data in Figure 3 make it clear that the secondary vesicles target the void regions during the backfilling step. Third, a striking feature of the images in Figure 3 is essentially the complete separation of TR and NBD emission, which was stable for several hours. This attribute of the image indicates that no noticeable diffusion of probe lipids occurs across the patterned features when fluid lipid bilayer patterns are backfilled with bilayers formed from cholesteroland sphingomyelin-rich SUVs. This result contrasts with the homogenization of the Texas red probe observed when the intercalating secondary phase was compositionally comparable to the primary pattern (Figure 2) and suggests kinetically arrested mixing.50,51 Taken together, these observations suggest that backfilling of patterned membranes is a simple and convenient approach for manipulating molecular compositions of supported bilayers and as a means of designing kinetically stabilized patterns of material heterogeneity within fluid-supported membranes. We note that these data do not conclusively establish the fusion of the intercalated vesicles into single bilayers. Later in the article, we show several independent lines of evidence based on fluorescence recovery and temperature and composition-dependent probe mixing to establish that the adsorption of secondary vesicles must indeed result in at least partial fusion into the bilayer to form substantial lateral connectivity presumably with a high density of defects due to gaps and unfused vesicles. We carried out a series of experiments to explore if both lo and ld phases of the phase-separating secondary bilayer remain kinetically trapped. The fluorophores used in the experiments reported above were phase-sensitive. Texas red-DHPE is known to have a high partition coefficient in the fluid phase (e.g., POPC), whereas NBD-DHPE partitions with some preference for the lo and gellike phases.52 Thus, the observed isolation of the fluorophore implies the dominant presence of fluid- or ld-phase molecules in the surrounding areas and the concentration of the lo phase within the backfilled region of the bilayer. Additional support for the concentration of lo-phase molecules in the “green” regions of the bilayer was obtained by examining the binding

of cholera toxin to monosialoganglioside (Gm1) molecules.53 Gm1 is a glycosphingolipid with a high affinity for association with chol and SM.54 In this regard, Gm1 reports on the cholesteroland SM-rich local environments within fluid membranes. It is also well known that CTB binds specifically to Gm1.55 Thus, using fluorescently labeled CTB, it is possible to probe any repartitioning of Gm1, and hence chol and SM, from the secondary phase to the primary phase. When the backfilled SUVs consisting of approximately equimolar POPC, SM, and cholesterol were doped with ∼2 mol % Gm1 molecules, no noticeable change in the arrested mixing of the primary and the secondary phases (see above) was observed. Epifluorescence images shown in Figure 4 reveal that FITC-CTB was dominantly localized in the square “green” regions of the sample (initial pattern voids which were subsequently backfilled with the secondary raft-forming bilayer mixtures), suggesting that Gm1 and thus chol and SM remain within the backfilled pattern elements. Next, to examine whether the fluid lipid molecules from the secondary mixture exchange with the surrounding bilayer phase, we switched the fluorophores. Specifically, Texas red-DHPE, which, as discussed above, has an affinity for the fluid phase, was deliberately used to dope domain-forming secondary SUVs. To enable fluorescence contrasts, NBD-DHPE was used to dope fluid POPC in our primary patterned bilayer. Epifluorescence images shown in Figure 5 unambiguously show that the TR-DHPE concentration is reversed from initial localization in secondary “square” elements (50) Jung, S. Y.; Holden, M. A.; Cremer, P. S.; Collier, C. P. ChemPhysChem 2005, 6, 423-426. (51) The line tension at the edge of the mixed composition bilayer can be expected to influence the shape of the intercalated lipid phase. In the present case, high cholesterol and sphingomyelin concentrations must impart some liquidordered or gellike character. We further note that the minimization of line tension in supported membranes was previously observed to be very slow, presumably because of frictional coupling with the substrate. (Kaizuka, Y.; Groves, J. T. Biophys J. 2004, 86, 905-912). (52) Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 1417-1428. (53) Holmgren, J.; Lonnroth, I.; Mansson, J. E.; Svennerholm, L. Proc. Natl. Acad. Sci. U.S.A. 1975, 72, 2520-2524. (54) Dietrich, C.; Volovyk, Z. N.; Levi, M.; Thompson, N. L.; Jacobson, K. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 10642-10647. (55) Kuziemko, G. M.; Stroh, M.; Stevens, R. C. Biochemistry 1996, 35, 63756384.

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Figure 5. Phase-dependent migration of the Texas red probe. Diffusion of Texas red-DHPE from the raft pattern to the POPC surroundings. Initial (a) green and (b) red channel epifluorescence images of a stamped bilayer pattern comprising an NBD-PE (3%)doped POPC bilayer pattern backfilled with TR-doped raft-forming SUVs (1% Texas red-DHPE, 39% POPC, 30% cholesterol, 28% sphingomyelin, and 2% GM1). (c) Green (d) red channel images after about 4 min. (See the text for details.)

to notably higher concentration in the surrounding grid areas. This rapid migration of Texas red-DHPE provides many important structural clues. First, it indicates that the molecules from the secondary phase can exchange with the primary phase, confirming at least a partial rupture to form sufficient lateral connectivity, rather than remain adsorbed as discrete unfused vesicles. This inference is further supported by fluidity measurements, the temperature and concentration dependence of the molecular exchange between the two phases, and their response to biochemical treatments (see below). Second, these results indicate that the fluid phase is concentrated in the grid area and that the Ch- and SM-enriched square elements are in the lo phase. Third, our observations are consistent with the picture (but do not directly establish) that the fluid PC molecules (e.g., POPC or DOPC) from the secondary phase equilibrate with the primary phase within the time scales of our experiments whereas cholesterolrich phase domains do not. Such a discrepancy in equilibration rates for the lo and ld phases appears reasonable because of large differences in the activation energy barriers for the mixing of supramolecular lo phase clusters than for ld phase molecules. Together, these two experiments suggest that secondary-phase intercalation results in nonequilibrium distributions of lo phase molecules whereas ld and fluid-phase molecules readily equilibrate, forming a single contiguous bilayer with a nonequilibrium spatial concentration of cholesterol-rich phases. Phase State of the Intercalated Bilayer. It is now well established that cholesterol interacts more strongly with SM than with unsaturated PC.20,21 It is also well understood that cholesterol has a condensing effect on the SM, resulting in a reduced effective molecular area. As a result, ternary mixtures comprising PC, SM, and chol are expected to exhibit liquid-liquid phase separation forming a chol- and SM-rich liquid-ordered or lo phase distinct from the liquid-disordered or ld phase of predominatly unsaturated PC. Indeed, significant evidence is now available that suggests that lipid mixtures comprising chol, SM, and PC mixtures phase separate into coexisting lo and ld phases for a broad range of molecular compositions15,22 and in a broad range of membrane configurations (e.g., lipid monolayers, bilayers derived from SUVs and Langmuir-Blodgett depositions, giant

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unilamellar vesicles, and multilamellar lipid stacks). Although there is a general consensus with regard to phase separation in these lipid mixtures, disagreements over the extent of phase separation and the sizes of microdomains remain.15 To probe the nature of the microphase separation of lo and ld phases in our intercalated bilayer patterns, we examined fluorescence emission in the raft-composition regions of our samples using high-resolution imaging (100×). The images shown in Figure 6 reveal that the NBD-PE emission within the secondary backfilled phase is nonuniform. The green regions of the images reveal a “grainy” fluorescence morphology covering the entire bilayer phase.56 Such nonuniformities in fluorescence may be due to the presence of some unfused vesicles or may indicate phase separation. Single-component fluid-supported membranes, such as those derived using POPC or DOPC SUVs, rarely exhibit such grainy fluorescence morphologies, supporting the notion that the observed composition-dependent fluorescence heterogeneity may be due to phase separation within the intercalated phase. Because NBD-PE52 has some preference for partitioning into the lo phase, the observed fluorescence heterogeneities in our data are consistent with the two-phase structure for the secondary green bilayer patterns: a raftlike lo phase wherein NBD preferentially resides and a nonraft ld phase with lower probe concentration. Further characterization of the morphology of the intercalated phase was difficult because of the limited resolution (1 to 2 µm) in our fluorescence measurements. We can, however, reliably conclude that phase separation, if any occurs, must be below the optical resolution in our experiments. Many previous studies of domain formation utilizing similar lipid compositions in supported membrane configurations provide a useful basis for morphological comparisons. Dietrich et al.52 used asymmetric single supported bilayers built by transfers of Langmuir films. The leaflet proximal to the glass substrate was either silanized monolayer or an egg-PC monolayer. They examined POPC (or DOPC)/cholesterol/sphingomyelin at 2(1): 1:1 molar concentrations. In all cases, they observed static, circular, or oval domains several micrometers (2-5 µm) in diameter using epifluorescence imaging. On the basis of partitioning characteristics of well-known probes (TR, FL, and NBD), they attributed these domains to cholesterol- and sphingomyelin-rich raftlike features. In a recent study, similar micrometer-scale stationary domains were reported by Stottrup et al.25 These large domains were shown to reflect the initial domain morphologies of the precursor Langmuir phase at the air-water interface. Such “quenching” of Langmuir phases has been previously interpreted in terms of kinetic barriers to reequilibration when condensed phases of Langmuir monolayers are transferred to glass substrates.57 In contrast, our results confirm that the surface spreading of small unilamellar vesicles does not show such large-scale domains. Indeed, Stottrup et al.25 also remarked that large domains were not observed when unilamellar vesicles such as those used in the present study were employed to form bilayers. Recently, several studies employing vesicle deposition procedures such as ours have reported the formation of submicroscopic-scale domain morphologies on mica substrates using scanning probe microscopy.30-33 Our results are entirely consistent with these reports but do not independently establish the detailed morphology of the lo and ld phase coexistence. (56) Note the presence of some extended structures in the fluorescence morphology. They were observed only in some bilayers formed by the rupture and spreading of SUVs derived using raft-forming lipid mixtures. We do not fully understand the origin of these structures but conjecture the deposition of occasional unfused vesicles. (57) Seul, M.; Subramaniam, S.; McConnell, H. M. J. Phys. Chem. 1985, 89, 3592-3595.

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Figure 6. Closer view of raftlike-engineered patterns. A green channel image of the phase-separating intercalated phase (3% NBD-PE, 45% POPC, 25% cholesterol, 25% sphingomyelin, and 2% GM1) obtained by backfilling within the 100 × 100 µm2 stamped POPC pattern. Oil-immersion objectives (10× and 100×) were used. (See the text for details.)

Figure 7. Fluidity patterns. Fluorescence recovery after photobleaching for the 50 × 50 µm2 membrane pattern comprising 1% Texas red-DHPE and 99% POPC in the grid region and the intercalated raft-forming lipid bilayer (33% DOPC, 31% SM, 31% chol, 2% GM1, and 3% NBDPE) in the square region. The dotted circle in b highlights the rapid blurring of the photobleached. (a-c) Images (60×) of before, immediately following, and 3 min after photobleaching TR. (d-f) Images (100×) of raft-enriched bilayer regions immediately following, 10 min after, and 51 min after photobleaching the NBD dye. (See the text for details.)

Another interesting attribute of the fluorescence heterogeneity within the intercalated green pattern elements in our samples is that it appeared immobile within the plane of the membrane on the time scales of our experiments (2-4 h). By contrast, raftlike heterogeneities in unsupported “free” bilayer membranes and GUVs exhibit lateral dynamics of phase-separated domains. To this end, previous studies suggest two plausible explanations. First, at the concentrations used in our study, the lo phase exceeds 45-50% of the membrane and constitutes the dominant phase, and the more fluid PC-rich ld represent the minority phase.58 In a previous atomic force microscopy study, Rinia and co-workers32 have shown that at comparable compositions (35:35:30 DOPC/ SPM/Ch) supported bilayers formed by vesicle fusion may phase separate to form a pattern of lo and ld phases wherein the lo phase becomes a percolating phase and thus remains immobile in the plane of the bilayer.59 Second, it has been variously suggested that the proximity of the substrate surface offers a drag for the mobilities of the lo microdomains, thus impeding their collective (58) Crane, J. M.; Tamm, L. K. Biophys. J. 2004, 86, 2965-2979. (59) Rinia and co-workers32 also observed that a small variation the composition (37.5:37.5:25 DOPC/SPM/Ch) changed the percolation characteristics, yielding densely packed, discrete nanometer-scale domains.

dynamics.60 The behavior of lipid mixtures employing a lower cholesterol concentration discussed later in the article offers further insights. Patterned-Fluidity-Supported Bilayers. We investigated whether our method of kinetically arresting very different local lipid environments within single bilayers results in patterns of distinct fluidities. We probed the long-range translational mobilities of the fluorescent probes with the TR-rich primary and the NBD-rich secondary phases of our patterned bilayers using FRAP. Selected frames from time-lapse FRAP measurements are shown in Figure 7. Using a simple adaptation of conventional microscopy-based FRAP, we observe that the TRrich surroundings and the NBD-rich square pattern elements exhibit near-complete recovery of bleached fluorophores. This confirms the essential fluidities and liquidlike character of the two juxtaposed phases. Furthermore, we observe that the recovery of the bleached TR fluorophore spot (∼25 µm in diameter) was quantitatively comparable to the recovery characteristics for preintercalated patterned and continuous bilayers (full recovery of a 35-µm-diameter spot within 4 to 5 min.) By contrast, the NBD-incorporating secondary phase(s) of the 1:1:1 DOPC/chol/ (60) Nissen, J.; Jacobs, K.; Radler, J. O. Phys. ReV. Lett. 2001, 86, 19041907.

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Figure 8. Epifluorescence images of pattern bluring by diffusion. (a and b) Initial fluorescence images of a stamped/backfilled bilayer comprising 3% NBD-PE, 47% POPC, 20% cholesterol, 28% sphingomyelin, and 2% GM1 as a backfilled phase within the stamped phase composed of 1% Texas red-DHPE and 99%POPC. (c-e) Red-channel images after 80, 120, and 196 min. (f) Green-channel image after 196 min.

SM mixture exhibit notably slower diffusion. (The complete fluorescence recovery requires ∼30-50 min depending on the size of the pattern element.) These measurements, in conjunction with a simple FRAP analysis (Experimental Section), yield a semiquantitative estimate of ∼1.5-2 µm2/s for the TR diffusion within the fluid POPC environment. Accurate quantitative information regarding the diffusion of NBD in raft-enriched square regions was difficult to obtain using the same samples because of the edge effects in confined secondary phases on probe diffusion. For the diffusion of NBD of companion samples, when the secondary raft-forming SUVs of identical composition were spread over the entire coverslip by vesicle fusion, the recovery time was 24 min (Supplemental Information) for a 90 µm FRAP spot, and an NBD-DOPC 96 µm FRAP spot was recovered in 13 min. These values result in probe diffusion constants of ∼1.3 µm2/s for raft-enriched bilayer regions and ∼3.2 µm2/s for the DOPC bilayer, in good general agreement with previous reports where the addition of comparable amounts of chol and SM was found to lower the DOPC diffusion coefficient by a factor of 2.58 The foregoing inferences of spatial patterns of fluidity and the percolation characteristics of the raftlike phase are also consistent with the experiment summarized in Figure 5. We recall that when Texas red-DHPE was incorporated within the raft-forming phase it exhibited a rapid and directional migration from the lo-phase-rich square pattern elements to the surrounding PC-rich fluid membrane phase (see above). Because Texas red-DHPE is known to have greater partitioning preference for the fluid phase, the observed migration of TR-DHPE from the lo-phase-enriched square patterns lends further support to the lateral contiguity and patterned fluidity picture for our bilayer patterns. It is important to note here that whereas TR-DHPE avoids the lo phase it can “permeate” through the lo phase.61 Thus, despite the percolating character of the lo phase, fluid-phase molecules can equilibrate with the surrounding primary POPC bilayer. Temperature, Degree of Saturation of PC Lipid, and Cholesterol Concentration Coordinate the Stability of Engineered Raftlike Heterogeneities in Supported Bilayers. The pattern stability was strongly influenced by the degree of (61) Kaizuka, Y.; Groves, J. T. Biophys. J. 2004, 86, 905-912.

unsaturation of phospholipid chain, the cholesterol concentration, and the sample temperature. We compared the patterns for lipid mixtures containing 10, 20, and 30 mol % cholesterol under ambient room-temperature conditions. Sphingomyelin was 28%, and the POPC fraction was varied accordingly. As seen in Figure 8, the samples containing less than 30 mol % cholesterol did not show stable isolation between the primary- and the secondaryphase lipid probes. The NBD-DHPE probe was observed to diffuse over time into the primary phase. Complete mixing occurred in ∼2 h when the cholesterol concentration in the secondary phase was 10% whereas ∼3 h was required for fluorescence homogenization when the cholesterol concentration was 20%. We further observed that by raising the sample temperature the time required for complete fluorescence homogenization for these samples could be considerably reduced. For instance, raising the sample temperature to ∼42 °C decreased the mixing time for the 10% cholesterol mixture above to 80 min, and 30% cholesterol concentration raftlike lipid mixtures revealed fluorescence homogenization in ∼10 min at 60 °C. These results are summarized in Table 1. Similarly, when the phospholipid was fully unsaturated (e.g., DOPC, Tm ) -20 °C), the raftlike membrane patterns for 1:1:1 DOPC/chol/SPM mixtures in POPC surroundings remained stable for ∼5 h. By contrast, POPC (Tm ) -2 °C)-based patterns under similar concentrations were stable for >24 h. This reduction in pattern stability for DOPC-based raftlike lipid mixtures can also be explained on the basis of the reduced overall immiscibility of the raft-enriched regions and fluid lipid surroundings because of greater fluidity of the DOPCbased lipid mixture. Taken together, these observations confirm that at lower cholesterol concentrations and at elevated temperatures the primary- and secondary-phase immiscibilities are lower, thus presenting lower-energy barriers to activate the mixing of the primary and secondary phases. It is useful to recall that the final concentrations upon mixing substantially reduce the effective cholesterol and SM concentrations and probably give rise to a homogeneous lipid phase. These data suggest that the interplay between temperature, PC unsaturation, and cholesterol concentration control the stability of the kinetically arrested heterogeneity patterns in our system.

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Figure 9. Detergent resistance of the raftlike secondary-phase mixture. Epifluorescence images before and after Triton X-100 treatments of the membrane pattern comprising the surrounding fluid bilayer (1% Texas red-DHPE and 99% POPC) and square patterns enriched in cholesterol-rich phases (3% NBD-PE, 37% POPC, 30% cholesterol, 28% sphingomyelin, and 2% GM1).

Figure 10. Cholesterol extraction epifluorescense images revealing fluorescence homogenization upon treatment with 10 mM methyl-βcyclodextrin for a stamped/backfilled membrane pattern comprising 1% Texas red-DHPE and 99% POPC (stamped) and raft-forming backfilled SUVs (3% NBD-PE, 57% POPC, 10% cholesterol, 28% sphingomyelin, and 2% GM1: (left) red/green-channel image and (right) green-channel image after 20 min of treatment. Table 1 conc of cholesterol in rafts 10% cholesterol rafts 20% cholesterol rafts 30% cholesterol rafts

stability of rafts at melting of rafts at room temperature, 22 °C higher temperature 2h 3.5 h ∞, observed for 4 days

80 min at 42 °C 115 min at 42 °C 10 min at 60 °C

Engineered Raftlike Patterns Retain Selected Properties of Biological and Biomimetic Rafts. The engineered raftlike patterns retain some properties of lipid rafts believed to exist in biological membranes. This assertion is based on two key classes of experiments described below. First, a common biochemical method of isolating many raftpartitioning proteins from cell membranes involves treatment with mild nonionic detergents such as Triton X-100 (triton).62 Although detergent treatment disrupts most lipid-lipid and lipidprotein interactions, chol- and SM-rich rafts63-65 resist solubilization, presumably because of their stronger intermolecular interactions in the lo phase. Indeed, resistance to extraction by detergent continues to serve as an operational definition of lipid rafts. To explore whether the raftlike patterns derived in our experiments offered comparable resistance to detergent solubilization, we treated our bilayers, comprising TR-DHPE-doped POPC patterns backfilled with a NBD-DHPE-labeled mixture of 1:1:1 mol/mol/mol POPC/chol/SM, with Triton solution (0.012% concentration) at room temperature and 2:1:1 (0.033% concentration) POPC/chol/SM at 4 °C. In both cases, we observe that within about 10 min the neighboring Texas red-POPC pattern (62) Schroeder, R. J.; Ahmed, S. N.; Zhu, Y. Z.; London, E.; Brown, D. A. J. Biol. Chem. 1998, 273, 1150-1157. (63) Schuck, S.; Honsho, M.; Ekroos, K.; Shevchenko, A.; Simons, K. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 5795-5800. (64) Cerneus, D. P.; Ueffing, E.; Posthuma, G.; Strous, G. J.; Vanderende, A. J. Biol. Chem. 1993, 268, 3150-3155. (65) Hanada, K.; Nishijima, M.; Akamatsu, Y.; Pagano, R. E. J. Biol. Chem. 1995, 270, 6254-6260.

was no longer visible in fluorescence, leaving behind the choland SM-containing NBD-rich phase, which remained immobile. Representative epifluorescence images are shown in Figure 9. It is useful to note here that the pattern elements in our samples most likely contain raftlike microdomains as a percolating phase and a nonraft fluid phase as a discrete phase. We could not determine whether the nonraft component was solubilized during the detergent treatment. Second, we explored the role of cholesterol in stabilizing the phase-separated patterns. For cholesterol extraction, we used methyl β-cyclodextrin, which is known to deplete cholesterol11,66,67 when incubated with living cells. MβCD is a carbohydrate molecule with a pocket for binding cholesterol and forming a water-soluble complex.6 When raftenriched patterns of bilayers comprising 10% cholesterol, 28% sphingomyelin, 57% POPC, 2% GM1, and 3% NBD-PE were treated with 10 mM methyl-β-cyclodextrin (MβCD), the initial fluorescence pattern was replaced by homogeneous fluorescence emission as shown in Figure 10. These observations are in good agreement with recent studies.44,54 and with the notion that MβCD induces concentration-dependent cholesterol extraction that subsequently results in a decrease in fluidity disparity between the pattern elements and the surrounding fluid phase.

Conclusions We have shown that stamping and backfilling provides a simple and convenient means to concentrate cholesterol- and sphingomyelin-rich raftlike membrane heterogeneities within predetermined regions of a contiguous fluid phospholipid bilayer. The secondary intercalants fuse selectively within the gaps of the primary bilayer pattern. The experimental evidence obtained in (66) Ohtani, Y.; Irie, T.; Uekama, K.; Fukunaga, K.; Pitha, J. Eur. J. Biochem. 1989, 28, 17-22. (67) Rao, M.; Peachman, K. K.; Alving, C. R.; Rothwell, S. W. Immunol. Cell Biol. 2003, 81, 415-423.

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this study suggests that the secondary intercalant develops sufficient connectivity to allow large-scale mixing and molecular redistributions. This likely involves at least the partial rupture and fusion of vesicles into a raft-enriched bilayer connected to the primary-phase bilayer. The rapid equilibration of the ld phase and arrested mixing of the lo phase in the raft-forming mixtures with the surrounding primary phase results in the formation of the contiguous fluid phase within which the cholesterol- and sphingomyelin-rich phase remain unequilibrated in predetermined spatial patterns. The stability of these pattern was observed to be directly dependent on temperature and the disparity in fluidity (and amount of cholesterol) of the primary stamped lipid phase and the secondary backfilled phase. These patterns of molecular composition correspond to patterns of fluidity and associated functions that should be exploitable for developing practical assays to probe raft-dependent membrane functions.

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Acknowledgment. Support for this work was provided by the U.S. Department of Energy under grant no. DE-FG0204ER46173 (A.N.P.) and The NSF Center for Biophotonics Science and Technology. The Center for Biophotonics, an NSF Science and Technology Center, is managed by the University of California, Davis, under cooperative agreement no. PHY 0120999. Photolithographically patterned silicon masters for soft lithography were prepared using microfabrication facilities at the Northern California Nanotechnology Center at UC Davis. Supporting Information Available: Epifluorescence images of NBD-PE/DOPC and NBD-PE/DOPC/cholesterol/sphingomyelin/GM1 bilayers. This material is available free of charge via the Internet at http://pubs.acs.org. LA052248D