Formation and Characterization of Phospholipid Monolayers

transitions within the LC. By doping the phospholipids with a fluorescently labeled lipid (Texas Red- ... proteins at cell membranes,1,2 the first sta...
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Formation and Characterization of Phospholipid Monolayers Spontaneously Assembled at Interfaces between Aqueous Phases and Thermotropic Liquid Crystals Jeffrey M. Brake, Maren K. Daschner, and Nicholas L. Abbott* Department of Chemical Engineering, University of WisconsinsMadison, 1415 Engineering Drive, Madison, Wisconsin 53706 Received July 13, 2004. In Final Form: November 23, 2004 This paper reports an experimental investigation of the self-assembly of phospholipids (L-R-phosphatidylcholine-β-oleoyl-γ-palmitoyl (L-POPC), dipalmitoyl phosphatidylcholine (DPPC), and L-R-dilauroyl phosphatidylcholine (L-DLPC)) at interfaces between aqueous phases and the nematic liquid crystal (LC) 4′-pentyl-4-cyanobiphenyl. Stable planar interfaces between the aqueous phases and LCs were created by hosting the LCs within gold grids (square pores with widths of 283 µm and depths of 20 µm). At these interfaces, the presence and lateral organization of the phospholipids leads to interface-driven orientational transitions within the LC. By doping the phospholipids with a fluorescently labeled lipid (Texas Red1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (TR-DPPE)), quantitative epifluorescence microscopy revealed the saturation coverage of phospholipid at the interface to be that of a monolayer with an areal density of ∼49 ( 8% relative to hydrated lipid bilayers. By adsorbing phospholipids to the aqueous-LC interface from either vesicles or mixed micelles of dodecyltrimethylammonium and phospholipid, control of the areal density of phospholipid from 42 ( 10 to 102 (18% of saturation monolayer coverage was demonstrated. Fluorescence recovery after photobleaching (FRAP) experiments performed by using laser scanning confocal microscopy (LSCM) revealed the lateral mobility of fluorescently labeled DPPE in L-DLPC assembled at the interface with the liquid crystal to be (6 ( 1) × 10-12 m2/s for densely packed monolayers. Variation of the surface coverage and composition of phospholipid led to changes in lateral diffusivity between (0.2 ( 0.1) × 10-12 and (15 ( 2) × 10-12 m2/s. We also observed the phospholipid-laden interface to be compartmentalized by the gold grid, thus allowing for the creation of patterned arrays of phospholipids at the LC-aqueous interface.

Introduction In vitro mimics of biological membranes are widely used to study and report events such as the binding of signaling proteins at cell membranes, 1,2 the first stages of the fusion of viruses with membranes,3,4 the entry of protein toxins into cells,5 and the action of a range of enzymes (such as phospholipases).2,6,7 For example, mixtures of phospholipids and proteins have been assembled at interfaces between aqueous phases and either air,8-17 oils,17-22 * To whom correspondence should be addressed. E-mail: abbott@ engr.wisc.edu. Fax: 608-262-5434. (1) Sozzani, S.; Agwu, D. E.; McCall, C. E.; O’Flaherty, J. T.; Schmitt, J. D.; Kent, J. D.; McPhail, L. C. J. Biol. Chem. 1992, 267, 20481. (2) Stahelin, R. V.; Cho, W. Biochem. J. 2001, 359, 679. (3) Charych, D. H.; Nagy, J. O.; Spevak, W.; Bednarski, M. D. Science 1993, 261, 585. (4) Conboy, J. C.; McReynolds, K. D.; Gervay-Hague, J.; Saavedra, S. S. J. Am. Chem. Soc. 2002, 124, 968. (5) Song, X.; Swanson, B. I. Anal. Chem. 1991, 63, 2097. (6) de Haas, G. H.; Bonsen, P. P. M.; Pieterson, W. A.; van Deenen, L. L. M. Biochim. Biophys. Acta 1971, 239, 252. (7) Bonsen, P. P. M.; de Haas, G. H.; Pieterson, W. A.; van Deenen, L. L. M. Biochim. Biophys. Acta 1972, 270, 364. (8) Lee, S.; Kim, D. H.; Needham, D. Langmuir 2001, 17, 5544. (9) Von Tscharner, V.; McConnell, H. M. Biophys. J. 1981, 36, 409. (10) Mansour, H.; Wang, D.-S.; Chen, C.-S.; Zografi, G. Langmuir 2001, 17, 6622. (11) Schindler, H. Biochim. Biophys. Acta 1979, 555, 316. (12) Pattus, F.; Desnuelle, P.; Verger, R. Biochim. Biophys. Acta 1978, 507, 62. (13) Denizot, B. A.; Tchoreloff, P. C.; Proust, J. E.; Puisieux, F.; Lindenbaum, A.; Dehan, M. J. Colloid Interface Sci. 1991, 143, 120. (14) Yang, B.; Matsumura, H.; Furusawa, K. Colloids Surf., B 1999, 14, 161. (15) Qui, R.; MacDonald, R. C. Biochim. Biophys. Acta 1994, 1191, 343.

hydrophilic solids,23-32 or hydrophobic solids.23-28,33-36 Measurements of interactions occurring at these lipidladen interfaces can be broadly characterized as either requiring labeling of the target molecules (e.g., with fluorescent labels)28,37 or requiring relatively (16) Wiedmann, T. S.; Cheng, S.-M. J. Colloid Interface Sci. 1991, 147, 531. (17) Adalsteinsson, T.; Yu, H. Langmuir 2000, 16, 9410. (18) Walker, R. A.; Conboy, J. C.; Richmond, G. L. Langmuir 1997, 13, 3070. (19) Thoma, M.; Muhwald, H. Colloids Surf., A 1995, 95, 193. (20) Walker, R. A.; Gruetzmacher, J. A.; Richmond, G. L. J. Am. Chem. Soc. 1998, 120, 6991. (21) Walker, R. A.; Gragson, D. E.; Richmond, G. L. Colloids Surf., A 1999, 154, 175. (22) Benda, A.; Benesˇ, M.; Marecˇek, V.; Lhotsky´, A.; Hermens, W. Th.; Hof, M. Langmuir 2003, 19, 4120. (23) Sackmann, E. Science 1996, 271, 43. (24) Tiberg, F.; Harwigsson, I.; Malmsten, M. Eur. Biophys. J. 2000, 29, 196. (25) Kalb, E.; Frey, S.; Tamm, L. K. Biochim. Biophys. Acta 1992, 1103, 307. (26) Elliott, J. T.; Burden, D. L.; Woodward, J. T.; Sehgal, A.; Douglas, J. F. Langmuir 2003, 19, 2275. (27) Keller, C. A.; Kasemo, B. Biophys. J. 1998, 75, 1397. (28) Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 1417. (29) Plant, A. L. Langmuir 1993, 9, 2764. (30) Rao, N. M.; Plant, A. L.; Silin, V.; Wight, S.; Hui, S. W. Biophys. J. 1997, 73, 3066. (31) Hubbard, J. B.; Silin, V.; Plant, A. L. Biophys. Chem. 1998, 75, 163. (32) Von Tscharner, V.; McConnell, H. M. Biophys. J. 1981, 36, 421. (33) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105. (34) Groves, J. T.; Ulman, N.; Boxer, S. G. Science 1997, 275, 651. (35) Groves, J. T.; Ulman, N.; Cremer, P. S.; Boxer, S. G. Langmuir 1998, 14, 3347. (36) Cremer, P. S.; Boxer, S. G. J. Phys. Chem. B 1999, 103, 2554.

10.1021/la0482397 CCC: $30.25 © 2005 American Chemical Society Published on Web 02/11/2005

Self-Assembly of Phospholipids

complex instrumentation (e.g., surface plasmon reflectometry).2,27,30,38 Recently, we described a new principle for the reporting of biomolecular interactions at lipid-laden interfaces.39 The approach revolves around the assembly of phospholipids at interfaces between aqueous phases and micrometer-thick films of thermotropic liquid crystals. We observed that the presence of the phospholipids at the interface of the liquid crystal (LC) led to well-defined orientations of the LC and that specific binding and enzymatic activity of proteins at these lipid-laden interfaces triggered orientational transitions in the LC. Whereas our past communication demonstrated the possibility of reporting biomolecular interactions at phospholipid-laden interfaces between thermotropic LCs and aqueous phases, this paper describes the results of a detailed investigation of methods that can be used to assemble phospholipids at these interfaces as well as some measurements of the properties of the resulting interfaces. We note that past studies have reported the transfer of phospholipids from aqueous dispersions of vesicles to interfaces to be a complex process that depends on a number of parameters, including the type of interface,21,27,35 vesicle size and concentration,11,12,15,21,31 phase state of the lipid,8,13,16,18 and solution conditions such as pH and ionic strength.13,14,25,36 In this paper, we report on the effect of the phase state and concentration of phospholipids (at constant pH and ionic strength) on the transfer of phospholipids to the interface of the LC. The resulting lipid-laden interfaces are characterized by the areal density of lipid (surface coverage relative to a bilayer), lateral heterogeneity, and lipid mobility. The results of this study provide guidelines for tailoring the properties of lipid-laden interfaces of LCs. Materials and Methods Materials. Dodecyltrimethylammonium bromide (DTAB), tris(hydroxymethyl)aminomethane (Tris), ethylenediaminetetraacetic acid (EDTA), calcium chloride, hydrochloric acid, chloroform, l-R-phosphatidylcholine-β-oleoyl-γ-palmitoyl (LPOPC), L-R-dipalmitoyl phosphatidylcholine (L-DPPC), D-Rdipalmitoyl phosphatidylcholine (D-DPPC), and L-R-dilauroyl phosphatidylcholine (L-DLPC) were obtained from Sigma-Aldrich (St. Louis, MO). Octadecyltrichlorosilane (OTS) and sodium chloride were obtained from Fisher Scientific (Pittsburgh, PA). Texas Red-1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (TR-DPPE) and N-(fluorescein-5-thiocarbamoyl)-1,2-dipalmitoylsn-glycero-3-phosphoethanolamine (F-DPPE) were purchased from Molecular Probes (Eugene, OR). All chemicals were used as obtained. Deionization of a distilled water source was performed using a Milli-Q system (Millipore, Bedford, MA) to give water with a resistivity of 18.2 MΩ cm. 4′-Pentyl-4cyanobiphenyl (5CB) was obtained from EM Sciences (New York, NY). The glass microscope slides were Fisher’s Finest Premium Grade obtained from Fisher. Gold specimen grids (20-µm thickness, 283-µm grid spacing, and 50-µm bar width) were obtained from Electon Microscopy Sciences (Fort Washington, PA). Sylgard 182 elastomer and curing agent (PDMS) were obtained from Dow Corning Corp. (Midland, MI), and Norland Optical Adhesive 61 (PU) was obtained from Norland Products, Inc. (Cranbury, NJ). Preparation of Optical Cells. Glass slides were cleaned in piranha solution (70% (v/v) sulfuric acid and 30% (v/v) hydrogen peroxide) for 1 h at ∼80 °C (warning: piranha solution reacts strongly with organic compounds and should be handled with extreme caution; do not store the solution in closed containers) (37) Yang, T.; Jung, S.-Y.; Mao, H.; Cremer, P. S. Anal. Chem. 2001, 73, 165. (38) Plant, A. L.; Brigham-Burke, M.; Petrella, E. C.; O’Shannessy, D. J. Anal. Biochem. 1995, 226, 342. (39) Brake, J. M.; Daschner, M. K.; Luk, Y.-Y., Abbott, N. L. Science 2003, 302, 2094.

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Figure 1. (A) Schematic illustration and (B) photograph of the experimental system used in this work. The scale bar in part B corresponds to 1 cm. according to published procedures.40 The glass slides were then either functionalized with OTS according to published procedures41 or partially coated with a thin film of gold. Gold was deposited onto the slides by physical vapor deposition using an electron beam evaporator (model VES-3000-C, TekVak Industries Inc., Brentwood, NY). An 8-nm-thick layer of titanium was first deposited to promote the adhesion of a 20-nm-thick film of gold to the glass slide. The deposition was performed at normal incidence.42 A second slide was used as a mask during the deposition process to yield a surface with an abrupt ( Tm; L-DLPC, Tm ) -1 °C; L-POPC, Tm ) -4 °C; D,L-DPPC, Tm ) 41 °C)8,43 resulted in a clear solution that was filtered twice using a 0.22-µm filter (Millipore). Quasi-elastic light scattering confirmed the presence of vesicles (diameter 36 ( 2 nm). The vesicles were typically used within 48 h of their preparation. We observed large (∼100 nm) aggregates forming after ∼1 week. Mixed micelles formed from phospholipids and DTAB were prepared by resuspending the dried lipid in TBS containing DTAB at a molar ratio of 30-300:1. The concentration of DTAB was kept constant at 3 mM, while the lipid concentration was varied. The presence of mixed micellar aggregates was confirmed by quasi-elastic light scattering (diameter 3.4 ( 0.2 nm). Formation of Phospholipid Layers. Lipid was deposited onto the surface of the LC within the optical cells by filling the polymer wells with ∼100-250 µL of either a dispersion of vesicles or a solution of mixed micelles of lipid and DTAB. Vesicle fusion was allowed to occur for 2 h to ensure complete adsorption of the lipid.11,12,14,15,21,25,27,31 Adsorption of phospholipids from mixed micelles was allowed to continue for 30 min, although typically no further change in the appearance of the system was observed after 5 min of equilibration. At the end of the equilibration period, the aqueous phase was flushed with ∼10 mL of TBS-Ca2+ that contained no lipid. Throughout the equilibration and rinsing procedures, the lipid-laden 5CB interface remained in contact with the aqueous phase. Patterned lipid layers were formed by first contacting a region of 5CB within the grid with a ∼100 nL droplet of TBS containing mixed micelles (30:1) of DTAB and L-DPPC doped with 1% F-DPPE for ∼30-60 s. The entire grid impregnated with 5CB was then immersed in ∼250 µL of TBS containing vesicles of L-DLPC and allowed to equilibrate for 2 h. Prior to imaging, the aqueous phase was flushed with ∼10 mL of TBS which contained no lipid. When studying the dynamics of fusion of vesicles with the aqueous-5CB interface, we used dispersions of vesicles that had been aged >3 weeks. These dispersions contained large vesicles (>95% of the vesicle population had a diameter of 100 ( 9 nm) that exhibited slower adsorption kinetics than freshly prepared vesicles.12 Two different methods were used to examine the adsorption of phospholipids to the aqueous-LC interface. (1) Vesicle dispersions were contacted with the 5CB for 2 h while imaging the 5CB with a polarized light microscope (see below). (2) Vesicle dispersions containing fluorescently labeled phospholipids were contacted with 5CB for a fixed time (typically between 15 and 120 min). The lipid-containing solutions were then washed out of the cell using ∼10 mL of TBS containing 5 mM calcium chloride (TBS-Ca2+). The rinsing step was required to remove the fluorescent lipid from the bulk solution. Polarized Light Microscopy. The orientation of 5CB was determined by using plane-polarized light with an Olympus BX60 microscope with crossed polarizers (transmission mode). Each optical cell was placed on a rotating stage located between the polarizers and imaged with the aqueous-5CB interface facing either up or down. The focus was adjusted to either the aqueous5CB interface (facing up) or the OTS-5CB interface (facing down). Identical optical textures in the 5CB were observed regardless of the orientation of the sample. Orthoscopic examinations were performed with the source light intensity set to 50% of full illumination and the aperture set to 10% in order to collimate the incident light. Homeotropic alignments were determined by first observing the absence of transmitted light during a 360° rotation of the sample. Insertion of a condenser below the stage and a Bertrand lens above the stage allowed conoscopic examination of the cell. An interference pattern consisting of two crossed isogyres indicated homeotropic alignment.44 In-plane birefringence was indicated by a bright, colored appearance of 5CB and the presence of brush textures, typically four-brush textures emanating from a line defect, when the sample was viewed between crossed polarizers.45 All images were captured using a (43) Marsh, D. CRC Handbook of Lipid Bilayers; CRC Press: Boca Raton, FL, 1990. (44) Bloss, F. D. An Introduction to the Methods of Optical Crystallograpy; Holt, Rinehart and Winston: New York, 1961. (45) Sonin, A. A. Freely Suspended Liquid Crystalline Films; John Wiley & Sons: New York, 1998.

Brake et al. digital camera (Olympus C-2040 Zoom) mounted on the microscope. The camera was set to an f-stop of 2.6 and a shutter speed of 1/320 s. Fluorescence Imaging of Phospholipid-Laden Aqueous-5CB Interfaces. Lipid layers comprised of 0.1-1% TRDPPE in L-DLPC or D-DPPC were formed at the aqueous-5CB interface according to the procedures described above. The adsorbed phospholipids were then imaged by epifluorescence microscopy using an Olympus IX71 inverted microscope equipped with a 100-W mercury lamp. A fluorescence filter cube with an excitation filter of 560 nm and an emission filter of 645 nm was used to visualize Texas Red fluorescence. Fluorescein-labeled phospholipids were monitored using a fluorescence filter cube with a 470-490-nm excitation filter and a 515-nm emission filter. Images were collected with a Hamamatsu 1394 ORCA-ER CCD camera (Bridgewater, NJ) connected to a computer and controlled through SimplePCI imaging software (Compix, Inc., Cranberry Twp., NJ). Throughout the imaging, the sample was oriented with the aqueous-5CB interface facing down (toward the objective) to allow concurrent focus of the aqueous-5CB interface and the grid surface. Quantification of the interfacial lipid concentration was performed with phospholipids doped with 0.1% TR-DPPE. Fluorescence imaging was performed using an objective power of 10× and an exposure time of 0.1 s. Images of the fluorescence background were also aquired under identical conditions in the absence of the TR-DPPE. The images were analyzed using SimplePCI software to obtain the average fluorescence intensity within uniformly bright regions of the lipid layer. Some regions of the lipid layers, typically near grid edges (or scratched regions of SiO2 and OTS surfaces, see below), showed abnormally bright (reflection from grid surface) or dark (incomplete bilayer/ monolayer formation) fluorescence intensities. These samples were omitted from further data analysis. The average fluorescence intensities of the lipid layers were corrected for the background values obtained in the absence of the TR-DPPE. Qualitative fluorescence recovery after photobleaching (FRAP) experiments (see below for quantitative measurements) were performed by focusing on a region of the lipid layer (typically the center of a grid square) using a 40× objective. The TR-DPPE within this region was then photobleached in a circular pattern by constant excitation until the emission fluorescence intensity reached a minimum (∼30-60 s). The recovery of fluorescence intensity within the bleached regions of the lipid layer was monitored for 2-24 h using a 4× objective and short exposure times (∼0.05-0.1 s) to minimize further bleaching. Diffusion Measurements by FRAP Using Laser Scanning Confocal Microscopy (LSCM). Lipid layers comprised of 1% F-DPPE in L-POPC, L-DLPC, or D-DPPC were formed according to the procedures described above. The lipid layers were then imaged using a BioRad 1024 laser scanning microscope equipped with a 15-mW mixed gas (Kr/Ar) laser and photomultiplier tube. The laser produces an excitation line at 488 nm, and the fluorescence emission is passed through a 522-nm short bandpass ((17 nm) filter, suitable for imaging F-DPPE. Prior to photobleaching, the lipid-laden interface was located by performing a z-section scan at low (3%) laser power until the maximum fluorescence intensity was observed. The region to be bleached was then located by focusing (10× objective at 10× zoom, 3% laser power) on the corner of a grid square. We also measured the mobility of phospholipids on SiO2. On these substrates, we bleached regions of lipid at boundaries between gold and SiO2 using gold patterned on the SiO2. The region of the lipid-laden interface was selectively photobleached by scanning the region at 100% laser power. After photobleaching, the recovery profile was imaged with the 10× objective at a reduced zoom of 3× (resolution of 0.627 µm/pixel) and a lower laser power (3-10%). Scans were performed at normal speed (∼1 s/scan) every 3-10 s until significant recovery of fluorescence within the bleached region was observed. During the recovery scans, additional photobleaching of F-DPPE was not observed. The images (512 × 512 pixels) obtained during the recovery scans were analyzed using SimplePCI software. At each time point, the image was cropped to an identical 40 × 255 pixel region centered on the step-edge bleaching front and located away from the corners at the boundary between bleached and unbleached

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lipid. The images were digitally translated into averaged gray scale fluorescence intensities along the bleaching profile. These profiles were fit to an error function shape with the curvature (A) and position of the step-edge (x0) as free parameters. For one-dimensional diffusion, the transient solution to lipid diffusion subject to a step-edge initial condition is (see Appendix I)46

cA(x,t) )

( )

x - x0 I(x,t) - Imed ) erf ) erf[A(x - x0)] (1) Imax - Imed x4Dt

where cA(x,t) represents a scaled (-1 to 1) F-DPPE concentration; I(x,t), Imed, and Imax represent the transient, median, and maximum fluorescence intensities, respectively; D is the diffusion coefficient of F-DPPE in the lipid layer; and A ) (4Dt)-0.5. By plotting 4A-2 as a function of recovery time for a given photobleaching experiment, the data was fit to a line passing through the origin with a slope equal to the diffusion coefficient. Additionally, the value of x0 was monitored as a function of time. Significant changes in x0 from its initial value were interpreted as deviation of the recovery profiles from the one-dimensional diffusion model proposed in eq 1. Significant variation in x0 was typically observed when A (expressed in units of inverse meters) was measured to be 2 h) with a dispersion of vesicles of L-DLPC in TBS at a bulk concentrations of lipid of 1 µM. (B) A cartoon representation of the boundary conditions within a film of 5CB located between the aqueous-5CB and OTS-5CB interfaces. (C) Optical texture of 5CB after 2 h of contact with a dispersion of vesicles of L-DLPC (100 µM) in TBS. The inset is an interference pattern obtained by conoscopic imaging of 5CB. (D) Cartoon representation of the uniform anchoring of the 5CB film after the adsorption of a monolayer of lipid at the aqueous-5CB interface. (E) Optical texture of 5CB after contact with a dispersion of vesicles of L-DLPC (100 µM) for 10 min. All images were obtained using crossed polars. Scale bar, 300 µm.

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Table 1. Fluorescence Intensity of Phospholipid-Laden Interfacesa lipid L-DLPC L-DLPC L-DLPC L-DLPC L-DLPC L-DLPC D-DPPC D-DPPC

aggregate state surface vesicles vesicles vesicles micelles (30:1) micelles (100:1) micelles (300:1) vesicles micelles (30:1)

SiO2 OTS 5CB 5CB 5CB 5CB 5CB 5CB

total surface fluorescence

relative lipid coverage

17 000 ( 200 8200 ( 1100 8200 ( 1200 8700 ( 1300 5400 ( 700 3600 ( 900 6100 ( 900 8000 ( 1600

1.00 ( 0.01 0.49 ( 0.06 0.48 ( 0.07 0.51 ( 0.08 0.32 ( 0.04 0.21 ( 0.05 0.36 ( 0.05 0.47 ( 0.09

a All fluorescence intensities were obtained using 0.1 mol % TRDPPE in the indicated phospholipids (L-DLPC or D-DPPC). The exposure time (0.1 s) and objective power (10×) were held constant throughout the acquisition of images. Baseline values of the total fluorescence intensity of bare SiO2/OTS in TBS (5000) and 5CB in TBS (5500) were subtracted from the total fluorescence intensities of the phospholipids on SiO2/OTS and 5CB, respectively. The ratios indicate the moles of DTAB to the moles of lipid.

faces,17,19,20,50 which, in the case of 5CB, appears to cause homeotropic anchoring.51 As reported previously, long-chain phospholipids (n > 8) are essentially insoluble in water.52 Therefore, once assemblies of phospholipids are formed at oil-water and solid-water interfaces, they remain irreversibly adsorbed even after exchange of the bulk aqueous solution with lipid-free aqueous solutions.36 We also observed the anchoring of 5CB after contact with a vesicle dispersion (100 µM) of L-DLPC to not change further upon the exchange of the aqueous phase containing lipid for one which did not contain lipid. The anchoring of 5CB remained homeotropic (optical texture identical to Figure 2C) for >1 week, consistent with the irreversible adsorption of L-DLPC at the aqueous-5CB interface. We next determined the extent of transfer of L-DLPC onto the aqueous interface of the LC. Although we predicted that the saturated interface would likely be laden with a monolayer of lipid, we note that past studies of the transfer of lipid onto interfaces from vesicles have revealed that subtle changes in the properties of the interface (e.g., Al2O3 versus SiO2) can lead to different interfacial assemblies.35 Here, we make use of past studies that have established that phospholipids can spontaneously assemble into bilayers on SiO2 surfaces24-26,34,35 and monolayers on silanized surfaces.24,32 We use the systems described in these past studies as reference states for quantitative fluorescence. By doping vesicles formed of L-DLPC with 0.1% TR-DPPE, we compared the relative fluorescence intensity of lipid layers formed at the aqueous-5CB interface with those transferred onto SiO2 and OTS surfaces (Table 1). We measured the fluorescence intensity of L-DLPC/TR-DPPE on OTS to be 49 ( 6% of the fluorescence intensity of L-DLPC/TR-DPPE on SiO2, consistent with the formation of a bilayer on SiO2 and a monolayer on OTS. A similar fluorescence intensity (48 ( 7%) was measured for layers of L-DLPC/TR-DPPE formed by vesicle fusion at the aqueous-5CB interface. These results, when combined, support our prediction that a monolayer of L-DLPC is formed at the aqueous-5CB interface upon contact of the interface with a dispersion of vesicles of L-DLPC. Whereas the studies described above deal with the equilibrium state of the lipid-saturated interface, we also (50) Jada, A.; Lang, J.; Candau, S. J.; Zana, R. Colloids Surf. 1989, 38, 251. (51) Barmentlo, M.; Vrehen, Q. H. F. Chem. Phys. Lett. 1993, 209, 347. (52) Deems, E. A. Anal. Biochem. 2000, 287, 1.

examined the spatial and temporal processes that accompany the transfer of lipid onto the interface of the LC. During the spontaneous assembly of phospholipids at the aqueous-5CB interface, we observed that transient dark domains (10-100 µm in size) in the LC appeared ∼10 min after contact of the LC with the aqueous dispersion of vesicles (0.1 mM L-DLPC; Figure 2E). The LC in the dark domains was found to be oriented homeotropically, while the surrounding LC remained in a near-planar orientation at the interface with the aqueous phase. We further examined the dynamics of the formation of lipid layers from vesicle dispersions of L-DLPC containing 1% TRDPPE (Figure 3). By adding the fluorescent probe to the lipid dispersions, it was possible to sequentially image the optical texture of the 5CB (using polarized light microscopy) and the lateral organization of the adsorbed lipid (as inferred from the fluorescent micrograph) using the same system (see below for additional discussion). Upon initial exposure to the dispersion of vesicles (1 week, confirming the compartmentalization of the grids and the patterning of the phospholipid compositions within the grids. We were able to quantify the mobility of phospholipids supported at an aqueous-5CB interface by performing

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Figure 6. Examples of photobleaching patterns formed by a confocal scanning laser microscope using (A) L-DLPC/F-DPPE monolayers supported at an aqueous-5CB interface and (B) L-POPC/F-DPPE bilayers supported on SiO2 substrates. The white boxes in parts A and B represent the sampling areas over which the fluorescence intensity profiles were collected. The axes shown in part A correspond to the labels used in part E. Scale bar, 50 µm. Fluorescence intensity (40 × 255 pixels) (C) 11 s and (D) 38 s after photobleaching of a monolayer of L-DLPC/ F-DPPE at the aqueous-5CB interface. (E) The fluorescence intensity data (O) from part C were plotted and fit to eq 1 (solid line). Equation 1 was also fitted (dashed line) to the fluorescence intensity data (9) obtained from part D.

FRAP experiments using LSCM.55,56 Initially, a square region at the corner of a grid square (Figure 6A) or at the edge of the gold-SiO2 boundary (Figure 6B) was photobleached. The recovery of fluorescence within the boxed regions was then followed over time (Figure 6C,D), and plots of the average fluorescence intensity along the x-direction were constructed (Figure 6E). Least-squares regression of the data using eq 1 allowed A and x0 to be evaluated as a function of recovery time (Figure 7). The data in Figure 7 demonstrate that a plot of 0.25A-2 versus recovery time (t) indeed follows a linear trend with a slope equal to the diffusion coefficient of the probe in the lipid layer (5 × 10-12 m2/s for F-DPPE in L-DLPC in Figure 7), and the position of the bleaching step-edge (x0) remained essentially unchanged throughout the analysis. A complete summary of these analyses is given in Table 2. To validate our methods, we also measured the fluidity of bilayers of L-POPC supported on SiO2. As seen in Table 2, we measured the diffusivity of F-DPPE in L-POPC to be (2 ( 1) × 10-12 m2/s. This value is in reasonable agreement with diffusivity measurements in L-POPC (55) Cheng, Y.; Prud’homme, R. K.; Thomas, J. L. Macromolecules 2002, 35, 8111. (56) Yuan, Y.; Velev, O. D.; Lenhoff, A. M. Langmuir 2003, 19, 3705.

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Table 2. Diffusivity of F-DPPE and NBD-PC at Various Interfaces with Aqueous Solutions lipid

probe

aggregate state

interface

diffusion coeff (10-12 m2/s)

L-POPC

F-DPPE TRITC-DPPE NBD-eggPE F-DPPE F-DPPE NBD-PC

vesicles vesicles vesicles vesicles vesicles chloroform spreading, area ∼0.45 nm2 micelles (30:1) micelles (100:1) micelles (300:1) vesicles micelles (30:1)

SiO2 SiO2 SiO2 5CB 5CB heptane

2(1 6(1 3.5 ( 0.5 6(1 6(1 20

5CB 5CB 5CB 5CB 5CB

6.0 ( 0.6 9(1 15 ( 2 0.2 ( 0.1 3.9 ( 0.4

L-POPC L-POPC L-POPC L-DLPC L-DLPC L-DLPC L-DLPC L-DLPC D-DPPC D-DPPC

F-DPPE F-DPPE F-DPPE F-DPPE F-DPPE

ref 26 25 17

viscosity µ1 and µ2 as a function of the reduced drag coefficient (ΛT())

DT )

kT 4πa(µ1 + µ2)ΛT()

(2)

where a is the characteristic radius of the cylinder, k is Boltzmann’s constant, and T is the temperature.60 The reduced drag coefficient can be determined numerically using methods reported by Hughes et al. for a particular interface and lipid system by determining the dimensionless constant () from the equation

)

Figure 7. Determination of the diffusion coefficient of F-DPPE in L-DLPC at an aqueous-5CB interface by linear regression (solid line) of 0.25A-2 versus t (b). The data follow a straight line with an intercept through the origin and a slope equal to the diffusion coefficient. The position of the step-edge (x0) obtained from fitting the fluorescence intensity data (0) is also shown.

using other probes and techniques (3.5 × 10-12 to 6 × 10-12 m2/s).25,26 We also measured the fluidity of monolayers of L-POPC and L-DLPC deposited at the aqueous-5CB interface from vesicle dispersions of the phospholipids. Both L-POPC (Tm ) -4 °C) and L-DLPC (Tm ) 1 °C) exist in the liquid crystalline state in vesicles and transfer to the interface as densely packed monolayers (see Table 1).8,18 The diffusion constant of F-DPPE in the resulting lipid-laden aqueous-5CB interfaces was measured to be (6 ( 1) × 10-12 m2/s for both phospholipids. By comparison, densely packed monolayers of DLPC at heptane-water interfaces give rise to higher lateral diffusivities of the probe lipid 1-acyl-2-[12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl]phosphatidylcholine (NBD-PC) (∼20 × 10-12 m2/ s).17 As discussed below, we speculate that the high lateral diffusivities measured at the heptane-water interface as compared to the aqueous-5CB interface may reflect the lower viscosity of heptane as compared to 5CB.57-59 Hughes et al. modeled the two-dimensional translational diffusion of cylinders (e.g., phospholipids) (DT) in membranes located at the interface between two fluids of (57) Lide, D. R. CRC Handbook of Chemistry and Physics, 74th ed.; CRC Press: Boca Raton, FL, 1993. (58) Sperkach, Y. V.; Sperkach, V. S.; Aliokhin, O. D.; Strybulevych, A. L.; Masuko, M. Mol. Cryst. Liq. Cryst. 2001, 366, 91. (59) Jadz˘ yn, J.; Dabrowski, R.; Lech, T.; Czechowski, G. J. Chem. Eng. Data 2001, 46, 110.

(

)

a µ1 + µ2 h η

(3)

where h is the length of the cylinder and η is the viscosity of the membrane.60 For phospholipids at the interface between water and 5CB, we choose a ∼ 4 Å, h ∼ 16 Å, η ∼ 0.1 Pa s,60 µ1 ∼ 0.001 Pa s,57 and µ2 ∼ 0.03 Pa s.58,59 The same parameters were used for phospholipids at an aqueous-heptane interface except for µ2, the viscosity of heptane, which was taken to be 0.0004 Pa s.57 Using these parameter values, we calculated the diffusivity of the probe in L-DLPC at the aqueous-heptane interface to be 2.1 times that at the aqueous-5CB interface. Our experiments yielded the value of the ratio to be 3.3. Discrepancies between predicted and measured values likely arise from simplifications present in the model. For example, the viscosity of 5CB varies significantly depending on its orientation relative to the shear force.61 We also note that different probe molecules were used in our experiments (F-DPPE) and the experiments reported at the heptaneaqueous interface (NBD-PC). Differences in probe properties may influence the measured diffusivities. In the studies of lipid mobility at heptane-water interfaces reported above, the authors demonstrated that, by controlling the surface concentration of the DLPC, the diffusivity of the probe lipid could be systematically varied.17 At high specific surface areas of DLPC (>0.70 nm2/molecule), they observed maximal lipid diffusivity. By decreasing the specific surface area below 0.70 nm2/ molecule, they measured a monotonic decrease in the diffusivity up to film collapse. We investigated the change in diffusivity of F-DPPE in L-DLPC at the aqueous-5CB interface with varying surface concentration of L-DLPC (achieved by using mixed DTAB-lipid micelles to deliver the lipid, see above). We note here that L-DLPC monolayers formed from mixed DTAB (3 mM)-L-DLPC (0.1 mM) micelles were found to possess identical diffusivities (6 × 10-12 m2/s) to those of monolayers of L-DLPC formed from (60) Hughes, B. D.; Pailthorpe, B. A.; White, L. R. J. Fluid Mech. 1981, 110, 349. (61) Belyaev, V. V. Phys.-Usp. 2001, 44, 255.

Self-Assembly of Phospholipids

Langmuir, Vol. 21, No. 6, 2005 2227

Figure 8. Distribution of F-DPPE in lipid monolayers of D-DPPC formed at the aqueous-5CB interface from (A and B) vesicles and (C) mixed DTAB-lipid micelles. All images shown were obtained by epifluorescence microscopy. Similar images were also obtained using LSCM. Scale bar, 50 µm.

vesicle dispersions. This result is consistent with our conclusion that similar surface concentrations of L-DLPC can be formed at the aqueous-5CB interface by adsorption from vesicle dispersion (∼48% relative to L-DLPC bilayers) and mixed surfactant-lipid micelles (∼51% relative to L-DLPC bilayers, see Table 1). By decreasing the relative coverage of lipid at the aqueous-LC interface by increasing the surfactant-to-lipid ratio at a fixed DTAB bulk concentration of 3 mM, we measured a significant increase in the lipid diffusivity ((9 ( 1) × 10-12 m2/s at ∼70% monolayer coverage and (15 ( 2) × 10-12 m2/s at ∼48% monolayer coverage, see Table 2). We made similar measurements of the diffusivity of F-DPPE in D-DPPC at the aqueous-5CB interface when the lipid monolayer was formed from either vesicle dispersions or mixed micelles. On the basis of the results above, we predicted that these two types of lipid-laden interfaces would have significantly different diffusivities due to the limited transfer of D-DPPC to the aqueous5CB interface from vesicles at ambient temperature. The diffusivity of the probe lipid in densely packed monolayers of D-DPPC (∼47% coverage relative to L-DLPC bilayers) formed by adsorption from mixed DTAB (3 mM)-lipid (0.1 mM) micelles was measured to be (3.9 ( 0.4) × 10-12 m2/s, slightly lower than the values measured using L-DLPC and L-POPC (∼(6 ( 1) × 10-12 m2/s). However, the D-DPPC monolayers (∼36% coverage relative to L-DLPC bilayers) formed using vesicles were measured to have lipid diffusivities that were significantly slower (∼(0.2 ( 0.1) × 10-12 m2/s) than those of the more densely packed monolayers prepared from mixed micelles. We speculate that the low apparent diffusivity of F-DPPE in D-DPPC layers formed by adsorption from vesicles may reflect the presence of lipid domains at the aqueous-LC interface. It has been shown that phase separation can result in the formation of fluid (D ∼ 10-12 m2/s) and condensed (D ∼ 10-14 m2/s) phases.62,63 An epifluorescence micrograph of a D-DPPC monolayer containing 1% F-DPPE formed by vesicle adsorption confirmed the presence of domains at the aqueous-LC interface. Initially, a heterogeneous distribution of FDPPE was observed (∼1-10 µm F-DPPE-rich domains, Figure 8A). After aging the interface for 24 h, the F-DPPErich domains were observed to ripen, increasing in size to ∼10-50 µm (Figure 8B). In contrast, densely packed monolayers of D-DPPC at the aqueous-5CB interface formed from mixed micelles were found to be homogeneous on the micrometer scale (Figure 8C). The homogeneous lipid layers were measured to possess high diffusivities ((3.9 ( 0.4) × 10-12 m2/s). We do not yet understand the (62) Korlach, J.; Schwille, P.; Webb, W. W.; Feignenson, G. W. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 8461. (63) Tanaka, K.; Manning, P. A.; Lau, V. K.; Yu, H. Langmuir 1999, 15, 600.

mechanism by which the heterogeneity of the interface decorated with D-DPPC slows the mobility of the probe lipid. We note here that the presence of the domains in the D-DPPC-laden interface did not lead to patterned orientations of the liquid crystals. In all cases, the appearance of the 5CB remained similar to Figure 3D. It is interesting to note that coexisting liquid expanded and liquid condensed domains of DPPC at air-water10 and oil-water19 interfaces adopt similar shapes, sizes, and distributions to those shown in Figure 8A,B. Further, the coexistence of liquid expanded and liquid condensed phases of DPPC has been reported to occur at surface coverages ranging from 68 to 83% of a densely packed monolayer at ambient temperatures.10,19 The surface coverage of D-DPPC monolayers formed from vesicles at the aqueous-5CB interface was found to be ∼72% of a densely packed monolayer. These results suggest that two coexisting phases of D-DPPC similar to the liquid expanded and liquid condensed phases may be forming at aqueous5CB interfaces by vesicle adsorption at T < Tm. Conclusion The main conclusions of this paper are threefold. First, by combining quantitative epifluorescence and polarized light microscopy, we have demonstrated methods that permit control over the areal densities of lipid assembled at the aqueous-LC interface. The methods revolve around the use of mixtures of soluble surfactants and phospholipids. The use of mixed surfactant-lipid micelles appears to be a general and facile method for controlling the interfacial lipid concentration. It is applicable to a range of phospholipids and is independent of the phase state of the pure lipid in bilayers. Second, the results reported in this manuscript establish a method based on FRAP and LSCM to measure the lateral diffusivity of phospholipids at the aqueous-5CB interface. Measurement of the lateral diffusivity of the phospholipids (∼(4-6) × 10-12 m2/s for densely packed monolayers) confirmed the fluidity of L-DLPC and L-POPC monolayers at the aqueous-LC interface. By controlling the method and type of lipid delivered to the aqueous-LC interface, the mobility of the phospholipids could be tuned from 0.2 to 15 × 10-12 m2/s. When D-DPPC was adsorbed to the interface from vesicles, the lateral diffusivity was low (∼(0.2 ( 0.1) × 10-12 m2/s) and domains were observed at the interface by using fluorescence microscopy. Third, lipid layers at the aqueous-LC interface were observed to be compartmentalized by the grid squares used to host the LC, thus permitting the formation of patterned arrays of phospholipids having different compositions among adjacent grid squares. In summary, the experimental systems and methods presented in this work advance the potential

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Brake et al.

utility of LCs as real-time amplifiers and transducers of molecular interactions within biomimetic films of phospholipids. Acknowledgment. This research was partially supported by the National Science Foundation through BES0330333 and the Center for Nanostructured Interfaces (NSF DMR 0079983). Support from the National Institutes of Health (NIH 5 T32 GM08349) is also acknowledged. The authors thank Lance Rodenkirch and the W. M. Keck Laboratory for Biological Imaging at the University of Wisconsin for assistance with the laser scanning confocal microscopy. Appendix I. Model of Patterned Photobleaching in One Dimension Diffusion of F-DPPE within lipid monolayers and bilayers supported on 5CB and SiO2, respectively, represents a transient, two-component, two-dimensional mass transport problem. In the absence of phospholipids adsorbing to or desorbing from the interface, the total molar flux of phospholipids at any point in the film is zero such that

∂nB ∂nC ∂nA )∂t ∂t ∂t

(A.4)

cA(x,t)|x)1 ) 1, for all t > 0

(A.5)

{

(A.6)

]

cA|t)0 )

0, x <  1, x > 

where  represents the position of the step-edge bleaching front. The solution to eq A.3 subject to eqs A.4-A.6 is46 ∞

cA(x,t) ) 1 +

∑

n)0

{ [( ) ] [ ]} Bn exp -

2n + 1 2π2 t 2 D cos

∂2cA ∂2cA ∂cA ) -D + 2 ∂t ∂x2 ∂y

(A.2)

(2n + 1)π x 2

(A.7)

where

For short times (typically 30-60 s for D ∼ the recovery profile away from the corners of the bleached and unbleached regions (see Figure 6A,B) of the lipid layer is not a function of y, thus reducing the partial differential equation in A.2 to a problem in x and t only 10-12

∂2cA ∂cA ) -D 2 ∂t ∂x

∂cA | ) 0, for all t > 0 ∂x x)0

(A.1)

where nA is the moles of fluorescently active F-DPPE, nB is the moles of photobleached F-DPPE, and nC is the moles of unlabeled lipid. We assume that the unlabeled lipid is uniformly distributed at the interface such that its net molar flux is zero. Therefore, Fick’s first law of diffusion reduces to64

[

When an infinite domain in x and a step-function initial condition (cA(x > x0, t ) 0) ) 1, cA(x < x0,t ) 0) ) -1) are assumed, the solution to eqn A.3 is an error function of the form shown in eq 1.46 To test our approximation of an infinite domain in x, we considered an alternate solution to eq A.3 based on a finite domain in x. By selecting x ) 0 such that this point represents the boundary between the lipid layer and the gold surface or gold grid, a different set of boundary conditions can be placed on eq A.3. On the basis of our observations and past studies,34,35,39 the gold surface arrests the diffusion of phospholipids such that the flux of lipid at the edge of the gold surface approaches zero. The x-position within the lipid layer was further scaled such that the fluorescence intensity remained at the maximum throughout the recovery process at x ) 1. By rescaling the fluorescence intensity data from 0 (no active F-DPPE) to 1 (100% active F-DPPE) over a finite domain in x, the boundary conditions and initial condition can be expressed as

cm2/s),

(A.3)

(64) Bird, R. B.; Stewart, W. E.; Lightfoot, E. N. Transport Phenomena; John Wiley & Sons: New York, 1960.

Bn )

[

]

(2n + 1)π -4  , n ) 0, 1, 2, ... (A.8) sin 2 (2n + 1)π

By fitting eqs A.7 and A.8 to the recovery profiles of F-DPPE in L-DLPC at an aqueous-5CB interface, the diffusion coefficient was calculated to be 5.4 × 10-12 m2/s which agreed with the value obtained using eq 1 (6 × 10-12 m2/s) within experimental error ((1 × 10-12 m2/s). Therefore, eq 1 was considered to be a valid description of the recovery profiles at short times. LA0482397