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Synthesis and Characterization of Supported Phospholipid Monolayers: A Correlative Investigation by Radiochemical Titration and Atomic Force Microscopy Theodore M. Winger† and Elliot L. Chaikof*,†,‡ School of Chemical Engineering, Georgia Institute of Technology, and Laboratory of Biomolecular Materials Research, Department of Surgery, Emory University School of Medicine, Atlanta, Georgia 30322 Received January 15, 1998. In Final Form: May 14, 1998 Membrane-mimetic lipid films with or without cholesterol were formed on alkylsilane self-assembled monolayers on glass by a process of lipid vesicle fusion. Using a combination of radiochemical titration with 14C-labeled dipalmitoylphosphatidylcholine (DPPC) and AFM imaging in an aqueous environment, we have characterized both molecular packing density and, in an homologous series of lipid monolayers, film thickness. These data confirm that a monolayer phospholipid membrane can be produced by vesicle fusion to an alkylated glass substrate.
Introduction It is recognized that the adverse events leading to the failure of many vascular prostheses are related to maladaptive biological reactions at blood- and tissuematerial interfaces. In response to these problems, and particularly thrombosis of the small caliber prosthesis, grafts have been coated either with albumin, heparin, or prostacyclin analogues which inhibit the clotting cascade and platelet reactivity or with relatively inert materials, such as poly(ethylene oxide).1,2 None of these approaches have been successful in a clinical setting. As an alternate strategy, we believe that the cell membrane is likely a more appropriate focal point for the design of biologically active surfaces because it both participates in molecular recognition processes and also serves as a template for the generation of more complex composite structures with other macromolecules. For example, in the past decade, membrane-mimetic systems have aided efforts aimed at understanding the mechanisms of blood coagulation at the sites of vascular wall injury and on artificial surfaces. In addition, substrate-supported membranes functionalized with proteins, peptide sequences, and carbohydrates have been used to evaluate biomolecular recognition and cell-surface interactions.3,4 Therefore, we believe that supramolecular membrane complexes on solid supports create useful systems for probing and potentially controlling clinically adverse processes on the surfaces of medical implants in a biological environment.5,6 Supported membrane-mimetic bilayers have been produced by direct adsorption of unilamellar lipid vesicles to * Corresponding author. Address: 1364 Clifton Road, N. E., Box M-11, Laboratory for Biomolecular Materials Research and Department of Surgery, Emory University, Atlanta, GA 30322. Phone: (404) 727-8413. Fax: (404) 727-3660. E-mail: echaikof@ surgery.eushc.org. † Georgia Institute of Technology. ‡ Emory University School of Medicine. (1) Eberhart, R. C.; Huo, H.-H.; Nelson, K. MRS Bull. 1991, 16, 50. (2) Chaikof, E. L.; Merrill, E. W.; Coleman, J. E.; Ramberg, K.; Connolly, R. J.; Callow, A. D. AIChE J. 1990, 36, 994. (3) Lawrence, M. B.; Springer, T. A. Cell 1991, 65, 859. (4) McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A. Biochim. Biophys. Acta 1986, 864, 95. (5) Chaikof, E. L. Chemtech 1996, 26, 17. (6) Winger, T. M.; Ludovice, P. J.; Chaikof, E. L. Biomaterials 1995, 16, 443.
hydrophilic substrates, such as glass or metallized surfaces;7 by transferring monolayers from an air-water interface using Langmuir-Blodgett (LB) or LangmuirSchaeffer techniques;8 or by a combination of both techniques in which the first leaflet of the bilayer is deposited on the substrate by an LB method followed by contacting the immobilized monolayer with a vesiclecontaining solution.9-13 Recently, supported lipid monolayers have also been produced by first assembling a layer of closely packed hydrocarbon chains covalently bound to the substrate followed by exposure to a suspension of either detergent/lipid micelles or unilamellar lipid vesicles.14-20 Typically, the alkylated support has been produced by the formation of a self-assembled monolayer (SAM) of alkanethiols or thiolipids on a gold-coated support. In this report, we have characterized the formation of a supported monolayer of phospholipids with or without cholesterol on a SAM of octadecyltricholorosilane (OTS) covalently assembled on borosilicate glass. Properties of the alkylated substrate and the supported lipid monolayer were characterized. The scientific motivation for our research was the development of a reproducible method for the preparation of stable, immobilized membranemimetic surfaces to be used in the characterization of celland protein-membrane interactions. (7) Salamon, Z.; Wang, Y.; Tollin, G.; Macleod, H. A. Biochim. Biophys. Acta 1994, 1195, 267. (8) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105. (9) Kalb, E.; Frey, S.; Tamm, L. K. Biochim. Biophys. Acta 1992, 1103, 307. (10) Wenzl, P.; Fringeli, M.; Goette, J.; Fringeli, U. Langmuir 1994, 10, 4253. (11) Scho¨pflin, M.; Fringelli, U. P.; Perlia, X. J. Am. Chem. Soc. 1987, 109, 2375. (12) Lang, H.; Duschl, C.; Gra¨tzel, M.; Vogel, H. Thin Solid Films 1992, 210/211, 818. (13) Frey, S.; Tamm, L. K. Biophys. J. 1991, 60, 922. (14) Plant, A. L.; Brigham-Burke, M.; Petrella, E. C.; O’Shannesay, D. J. Anal. Biochem. 1995, 226, 342. (15) Plant, A. L. Langmuir 1993, 9, 2764. (16) Spinke, J.; Yang, J.; Wolf, H.; Liley, M.; Ringsdorf, H.; Knoll, W. Biophys. J. 1992, 63, 1667. (17) Seifert, K.; Fendler, K.; Bamberg, E. Biophys. J. 1993, 64, 384. (18) Heyse, S.; Vogel, H.; Sa¨nger, M.; Sigrist, H. Prot. Sci. 1995, 4, 2532. (19) Florin, E. L.; Gaub, H. E. Biophys. J. 1993, 64, 375. (20) Duschl, C.; Liley, M.; Corradin, G.; Vogel, H. Biophys. J. 1994, 67, 1229.
S0743-7463(98)00077-8 CCC: $15.00 © 1998 American Chemical Society Published on Web 06/24/1998
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Experimental Methods Materials. Dilaurylphosphatidylcholine (DLPC), dimyristoylphosphatidylcholine (DMPC), and DPPC were obtained from Avanti Polar Lipids (Alabaster, AL). Multi-Terge, an alkaline chelating detergent, was purchased from E. M. Diagnostic Systems (Gibbstown, NJ). 1,2-Di(1-14C)palmitoyl-L-3-phosphatidylcholine (14C-DPPC, 99%) was obtained from Amersham Life Science (Arlington Heights, IL) with a specific activity of 112 mCi/mmol. ScintiSafe Plus 50% scintillation fluid was from Fisher Scientific. Microscope borosilicate glass cover slips (S/P Cover Glass, 24 mm × 40 mm × 0.21 mm) were purchased from Baxter Scientific, and all other chemicals and solvents (HPLCgrade) were from Aldrich Chemicals. Dust-Off puff-duster cans (compressed gas filtered to 0.1 µm) were purchased from Falcon Safety Products, Inc. (Branchburg, NJ). Distilled water was deionized and of ultrafiltered grade (18 MΩ‚m, Continental). Preparation of Alkylated Glass. Cover slips were puffdusted and transferred to a Class 100 clean room. A solution of Multi-Terge/deionized water 1/8 v/v was applied, and the slides were rinsed with water and chloroform. Surfaces were blown dry with a nitrogen gun and exposed to an argon plasma (500 ( 10 mTorr) for 9 min at 100 ( 1 W. Slides were immediately immersed in an octadecyltrichlorosilane (OTS) reaction mixture for times ranging from 5 to 1320 min. The latter was prepared by mixing 248 mL of bicyclohexyl, 24.8 mL of hydrated CHCl3, and 27 mL of a solution of 63 mM OTS in anhydrous CCl4 (95%, stored in a desiccator over P2O5 at room temperature). After a defined incubation period, the slides were rinsed with chloroform, sonicated (∼47 kHz, 80 W) for 10 min in chloroform, rinsed with water, and dried with nitrogen.21 Vesicle Formation. Vesicles were produced by freeze-thaw cycling and subsequent extrusion through 600 and 200 nm polycarbonate membranes. The liposome extruder was built inhouse on the basis of the design of a similar device by MacDonald et al.22 Polycarbonate extrusion membranes (mean pore sizes: 100-600 nm) were obtained from Fisher Scientific and cut to size to fit the extruder. Briefly, a 10 mM lipid suspension (20 mM sodium phosphate at pH 6.2) was extruded 21 times through a double 600 nm porosity polycarbonate membrane, followed by another 21 extrusions through a 200 nm membrane. To ease the vesicle breakup and reassembly, each mixture was extruded at a temperature about 10-20 °C above the melting temperature of the given phospholipid. To ensure vesicle size at fusion, the liposome suspension was used within a few hours in subsequent fusion experiments. For radiochemical titration studies, vesicles were doped with 3 mol % 14C-DPPC. Formation of a Supported Membrane. Alkylated glass surfaces were incubated with a vesicle solution (150-200 µM) in phosphate buffered saline (PBS; 20 mM sodium phosphate, 150 mM NaCl) at pH 7.4. Fusion was conducted at 50 °C for DPPC-based systems and at room temperature (23 °C) for DMPCand DLPC-containing vesicles. After a defined incubation period, surfaces were rinsed copiously by repeated exchanges with PBS at room temperature. A 5 min contact time was allowed for each rinse, which afforded a 3-fold dilution of the previous solution. For radiochemical titration experiments an aluminum chamber was designed to hold the alkylated slides with wells of defined dimension (3.3 mm i.d.). All fusion experiments were performed in a humidity-saturated environment. Contact Angle Goniometry. Advancing and receding contact angles were measured on a Rame-Hart goniometer, Model 100-00. The values reported are an average of at least five readings on four separate samples. Radiochemical Titration Analysis. 14C-labeled samples were placed in ScintiSafe Plus 50% scintillation fluid (Fisher Scientific) and counted for 1 min using a 1211 Rackbeta liquid scintillation counter (LKB Instruments Inc., Gaithersburg, MD). Atomic Force Microscopy. All images were obtained using a Nanoscope IIIA instrument from Digital Instruments (Santa Barbara, CA) fitted with either a J-scanner or an E-scanner. Contact AFM cantilevers with the nominal spring constant 0.12 (21) Calistri-Yeh, M.; Kramer, E. J.; Sharma, R.; Zhao, W.; Rafailovich, M. H.; Sokolov, J.; Brock, J. D. Langmuir 1996, 12, 2747. (22) MacDonald, R. C.; MacDonald, R. I.; Menco, B. P. M.; Takashita, K.; Subbarao, N. K.; Hu, L.-R. Biochem. Biophys. Acta 1991, 1061, 297.
Langmuir, Vol. 14, No. 15, 1998 4149 N/m and featuring a pyramidal Si3N4 oxide-sharpened tip were purchased from Park Scientific Instruments (Sunnyvale, CA). Probe forces were typically 0.5-1 nN. Images in air (OTS/glass) were acquired at constant height in contact or tapping modes at scan rates of 4 and 1 Hz, respectively. Characteristically, three different areas, separated by at least 10 µm, were imaged. The average roughness (Ra) was calculated using the system software and is the average of triplicate measurements. To enable AFM imaging in water of fused phospholipid monolayers, a sample holder was designed for the piezoelectric stage which accommodated vesicle fusion, rinsing, and image analysis without the need for sample transfer, thus avoiding the risk of SAM exposure to air. Circular hydrophilic wells (1 mm deep, 8 mm i.d., 10 mm o.d.) were produced on a microscope slide by injection molding using medical grade silicon rubber (Technical Products Inc., Decatur, GA) and a 16 well chamber slide template (Nunc, Naperville, IL) followed by a 7 day cure at 37 °C in a humid environment. The wells were separated from each other with a glass cutter, and a 4 mm × 4 mm OTS/glass sample was glued to the bottom of each well with silicone rubber. After completion of the cure, these wells were used for vesicle fusion. All AFM samples of supported lipid monolayers were prepared in duplicate following the vesicle fusion and rinsing protocols described above. Images were acquired in an immersed contact regime, using the constant height mode, with a horizontal resolution set at 512 pixels × 512 pixels and a scanning line speed of 4 Hz. The probe force was maintained at 0.5-1 nN throughout the imaging experiment. As previously noted, three different areas, separated by at least 10 µm, were imaged. Electron Spectroscopy for Chemical Analysis (ESCA). Angle-dependent ESCA was performed on selected surfaces using a Physical Electronics (PHI) model 5100 spectrometer equipped with a Mg/Ti dual-anode source and an Al/Be window. The system used a hemispherical analyzer with a single-channel detector. Mg KR X-rays (1253.6 eV) were used as an achromatic source, operated at 300 W (15 kV and 20 mA). The base pressure of the system was lower than 5 × 10-9 Torr, with an operating pressure up to 10-7 Torr. A pass energy of 89.45 eV was used for acquisition of the survey spectra, and a pass energy of 35.75 eV was used for the high-resolution spectra of elemental regions. Spectra were obtained at takeoff angles of 15, 45, and 90°. The instrument was calibrated with Mg KR x-radiation by setting the distance between Au 4f7/2 and Cu 2p3/2 to 848.67 eV. The work function was set using Au 4f7/2 and Cu 2p3/2 and was checked with Au 3d5/2. All metals were sputter-cleaned to remove oxides. The full-width at half-maximum for Ag 3d3/2 was measured to be 0.8 eV for 30 000 counts.
Results and Discussion Preparation of a Self-Assembled Monolayer of OTS on Borosilicate Glass. Borosilicate glass cover slips were incubated with an OTS solution and removed from the reaction mixture at predefined time points for surface analysis by contact angle goniometry and AFM. Advancing contact angles increased from 28.3 ( 4.9° to 103.3 ( 1.5° after 20 min of static deposition and plateaued at 111.3 ( 1.0° after 50 min (Figure 1). Similar findings have been observed by other investigators with reported contact angles ranging between 108°23-27 and 112°.21,28-30 The receding contact angle of the fully formed OTS monolayer was 109 ( 2°. Prior to surface treatment, AFM analysis revealed that borosilicate glass was uniformly (23) Silberzan, P.; Le´ger, L.; Ausserre´, D.; Benattar, J. L. Langmuir 1991, 7, 1647. (24) Peach, S.; Polak, R. D.; Franck, C. Langmuir 1996, 12, 6053. (25) Nakagawa, T.; Ogawa, K. Langmuir 1994, 10, 525. (26) Angst, D. L. Langmuir 1991, 7, 2236. (27) Brzoska, J. B.; Azouz, I. B.; Rondelez, F. Langmuir 1994, 10, 4367. (28) Wasserman, S. R.; Whitesides, G. M.; Tidswell, I. M.; Ocko, B. M.; Persham, P. S.; Axe, J. D. J. Am. Chem. Soc. 1989, 111, 5852. (29) Tillman, N.; Ulman, A.; Schildkraut, J. S.; Penner, T. L. J. Am. Chem. Soc. 1988, 110, 6136. (30) Tidswell, I. M.; Rabedeau, T. A.; Pershan, P. S.; Folkers, J. P.; Baker, M. V.; Whitesides, G. M. Phys. Rev. B 1991, 44, 10869.
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Figure 1. Borosilicate glass cover slips were incubated with an OTS solution, removed from the reaction mixture at predefined time points, and analyzed by contact angle goniometry. Data are presented as average advancing contact angle with the inset providing additional detail with respect to early observation time points (20-120 min.).
smooth, with an average roughness of 1.1 Å over 1 × 1 µm2 (Figure 2). After 20 min of incubation in an OTS solution, surface “islands” were present and covered approximately 50% of the total area. At 35 min, the OTS layer covered 95% of the surface, which featured welldefined holes of an average diameter of 40 nm. The surface was free of defects after 60 min with an Ra of 1.5 Å. Observed surface features were similar whether obtained in contact or tapping mode. Conflicting reports have appeared regarding the mechanism of OTS monolayer formation. The observation of large surface domains in partial monolayers has lead some groups to speculate that alkyltrichlorosilanes form cross-linked networks in the solution before bonding to the hydroxylated substrate.31,32 Others have failed to observe significant domain formation and have suggested that silanes form a self-assembled monolayer by binding randomly to the surface.24,28,30 Reaction conditions including surface adsorbed water, temperature, and the spacing and density of reactive surface hydroxyl groups all likely influence the molecular pathway to monolayer formation.31,33 Our observation of island growth on borosilicate glass with subsequent “infilling” of holes does not support recent friction force microscopy studies by Peach et al.,24 who failed to observe the formation of significant OTS domains on a similar substrate. These investigators have suggested that their results could be attributed to the irregular spacing of hydroxyl groups on amorphous, noncrystalline glass surfaces. In contrast, our findings are consistent with those of Banga et al.,34 who noted island formation in the generation of an OTS SAM on industrial float glass. The protocol used in this report reproducibly afforded smooth alkylated glass samples of high quality, even upon scaleup to large batches. Angle-dependent ESCA provided additional evidence that a self-assembled OTS monolayer was formed (Figure (31) Flinn, D. H.; Guzonas, D. A.; Yoon, R.-H. Colloids Surf., A 1994, 87, 163. (32) Bourdieu, L.; Maaloum, M.; Silberzan, P.; Ausserre, D.; Coulon, G.; Chatenay, D. Ann. Chim. (Paris) 1992, 17, 229. (33) Parikh, A. N.; Allara, D. L.; Azouz, I. B.; Rondelez, F. J. Phys. Chem. 1994, 98, 7577. (34) Banga, R.; Yarwood, J.; Morgan, A. M.; Evans, B.; Kells, J. Langmuir 1995, 11, 4393.
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3). At takeoff angles of 15, 45, and 90°, mean free path estimations yield sampling depths of 20, 55, and 100 Å, respectively.35 On the basis of these estimates, we measured a monolayer thickness of approximately 30 Å, which agrees with an expected thickness of 23-27 Å26,28 for a monolayer with an alkyl chain tilt angle between 0° and 31°.29,36 Preparation of a Supported Phospholipid Monolayer: Analysis of Molecular Packing by Radiochemical Titration. Following lipid vesicle fusion (DPPC doped with 3 mol % 14C-DPPC) with OTS/glass substrates, samples were rinsed and the amount of 14CDPPC removed from the sample was measured for each PBS rinse. Twelve to twenty-four buffer exchanges were required to remove all residual radioactivity from the fusion wells (Figure 4). Slide 12 illustrates that a final organic rinse with chloroform/methanol 2/1 v/v removed residual surface-bound radioactivity, indicative of an adsorbed phospholipid layer which is resistant to further PBS rinses. The presence of an unaltered underlying OTS monolayer was confirmed by goniometry, which demonstrated advancing and receding water contact angles of 111° and 109°, respectively. Therefore, hydrophobic coupling between the two monolayers is strong and not reversible by extended wash procedures, even in the probable absence of significant alkyl chain interdigitation. After completion of rinses 6, 12, 18, and 24, the glass slide was removed and surface associated radioactivity measured. Given a known exposure area, the molar ratio of 14C-DPPC/DPPC (3.3:96.7), the specific activity of the radiolabeled probe (112 mCi/mmol), and the counting efficiency of the scintillation counter, we determined average molecular areas for DPPC in the monolayer ranging between 72 and 91 Å2/molecule (Figure 5). The temperature 50 °C was chosen during the adsorption of DPPC to the OTS monolayer in order to perform vesicle fusion above the main transition temperature of DPPC (41.5-42 °C). Under these conditions DPPC was in a liquid-expanded phase which is characteristically associated with a molecular area of about 60-70 Å2.37 For example, in compressibility studies using dilute multilamellar vesicles, Lis et al.38,39 observed a molecular area of 71.2 Å2 for DPPC at 50 °C in plain water. While this is consistent with our observations, it is likely that molecular packing of the phospholipid monolayer on the OTS/glass substrate varied with temperature. Following vesicle fusion and subsequent cooling to room temperature, DPPC passed through its main transition with a probable reduction of the cross-sectional area of its hydrocarbon chains. For example, Wenzl et al.10 measured a molecular area for DPPC of 36.5-45.1 Å2 at 25 °C after deposition on a DPPA monolayer. As such, we would predict that a reduction in PC molecular area would lead in our system to the development of lateral film tensionscompensated by either a corresponding hydrocarbon chain tilt or the development of surface defects in the lipid film. Although issues of molecular orientation and spatial organization could not be addressed by radiochemical titration, this methodology unequivocally confirmed the formation of a supported lipid monolayer. Preparation of a Supported Phospholipid Mono(35) Bierbaum, K.; Kinzler, M.; Wo¨ll, C.; Grunze, M.; Ha¨hner, G.; Heid, S.; Effenberger, F. Langmuir 1995, 11, 512. (36) Ulman, A. Chem. Rev. 1996, 96, 1533. (37) Hauser, H.; Pascher, I.; Pearson, R. H.; Sundell, S. Biochim. Biophys. Acta 1981, 650, 21. (38) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 657. (39) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 667.
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Figure 2. AFM analysis of alkylsilane monolayer formation on a borosilicate glass substrate as a function of incubation period in an OTS solution. Depicted time points: 0 (A), 20 (B), 35 (C), and 60 (D) min.
Figure 3. Angle-dependent ESCA to characterize the atomic percent surface composition of a fully formed self-assembled OTS monolayer. Sampling depth corresponds to takeoff angles of 15°, 45°, and 90°.
layer: AFM Imaging of Membrane-Mimetic Lipid Surfaces. To investigate film morphology, AFM in
Figure 4. Removal of 14C-DPPC with PBS. For slide 12, rinses 23 and 24 consisted of chloroform/methanol (2:1 v/v) and revealed the presence of a surface-bound phospholipid layer.
contact mode was performed under PBS and at room temperature on a series of surfaces composed of an homologous set of saturated phosphatidylcholines DLPC, DMPC, and DPPC. The DPPC surface appeared smooth
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Figure 5. DPPC packing density per unit area determined after completion of rinses 6, 12, 18, and 24. Average molecular areas for DPPC in the monolayer ranged between 72 and 91 Å2/molecule.
with an average roughness (Ra) of 1.4 Å measured over a 10 µm × 10 µm area (Figure 6). Holes of a deconvoluted diameter of about 50 nm and an average depth of 20 Å were observed and covered less than 1% of the total surface area examined. DMPC surfaces exhibited similar roughness values, but surface holes were not observed. Surfaces of DLPC exhibited infrequent holes with an apparent diameter of 60 nm and a depth of 13 Å. Hui et al.40 have reported similar surface defects in AFM studies of DPPE monolayers on a DPPE-on-mica substrate with the detection of bilayer-spanning holes with a depth of 54 ( 1 Å. In this study, both leaflets of the deposited lipid bilayer separated, exposing the hydrophilic mica surface. The observation of surface hole formation in our report suggests that small defects form in the supported monolayer despite the energetically unfavorable nature of exposing the underlying hydrophobic substrate to an aqueous environment. Therefore, while lipid molecules undergo chain tilting in order to maintain close-packed contact, the maximum tilt angle is insufficient to enable complete OTS surface coverage. Interestingly, the 7 Å difference in monolayer depth which was noted between DPPC and DLPC monolayers is consistent with a drop of 6 Å predicted from simplified molecular models of alkyl chain lengths.41 In defining this structural model, we note that monolayer thickness is comparable to measurements obtained from other model lipid membrane systems including freely suspended bilayers, vesicle bilayer dispersions, and LB monolayers on solid supports. Typical bilayer thickness characterized using a variety of physiochemical methods, including neutron and X-ray diffraction techniques, ranges between 60 and 70 Å.8,42 These measurements include solvent effects which probably account, at least in part, for the disparity in thickness measurements between these systems and those obtained from a supported lipid monolayer.7,43,44 For example, in recent reports of C16 (40) Hui, S. W.; Viswanathan, R.; Zasadzinski, J. A.; Israelachvili, J. N. Biophys. J. 1995, 68, 171. (41) Bain, C. D.; Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides, G. M.; Nuzzo, R. G. J. Am. Chem. Soc. 1989, 111, 321. (42) McIntosh, T. J. Biophys. J. 1980, 29, 237. (43) Nagle, J. F.; Wiener, M. C. Biochim. Biophys. Acta 1988, 942, 1. (44) Tien, H. T.; Diana, A. L. In Chemistry and Physics of Lipids; Bergelson, L. D., Chapman, D., Finean, J. B., Mangold, H. K., Shapiro, D., De Hass, G. H., Eds.; North-Holland: Amsterdam, 1968; pp 55101.
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lipid monolayers which were deposited on an alkanethiol SAM, film thicknesses of approximately 25 Å were measured by surface plasmon resonance spectroscopy.19,45 Vesicles were also prepared with a 33% molar content of cholesterol (Chol), fused to OTS glass, and imaged by AFM. DPPC/Chol monolayers exhibited more irregular surface morphology than DPPC alone, with a mean roughness of 1.9 Å over 5 × 5 µm2. Occasional surface mounds of about 150-200 nm in diameter were also observed (Figure 7). Supported DMPC/Chol and DLPC/ Chol surfaces also revealed the presence of surface “bumps”. The dimensions of these features in all cases were consistent with those of adsorbed liposomes. Surface holes were not detected in any of the cholesterol-containing films. AFM scratch tests were performed on PC/Chol monolayers above 10 nN, the known scratch resistance threshold for phospholipid bilayers on mica.40,46 An expected furrow was not observed, perhaps due to healing of the induced defect by rapid lateral diffusion of the vicinal phospholipids. In support of this interpretation, we calculated that the rate of monolayer removal from the surface was 0.16 µm2/s, assuming an AFM tip radius of 20 nm. This is significantly slower than the measured lateral diffusion coefficient for saturated PCs in PC/Chol bilayers, which is on the order of 10-8 cm2/s or 1 µm2/s.47 While this is an appealing explanation, reported lateral diffusion constants for supported lipid monolayers are characteristically 10-11 to 10-12 cm2/s, which would be inconsistent with our observations.48 We suspect that the healing of the furrow is accelerated by the unfavorable increase in surface energy when the underlying hydrophobic OTS layer is exposed to the aqueous environment. In summary, data provided by both radiotitration and AFM analyses are consistent with the formation of a supported phospholipid monolayer on an OTS SAM following a 16 h incubation period. While LangmuirBlodgett techniques have been utilized extensively for the production of supported lipid films, we believe that vesicle fusion has an important advantage in that this methodology, in principle, can be extended to a variety of alkylated surface geometries including nonplanar surfaces, as well as porous substrates. Typically, surface fusion of phospholipid vesicles is initiated by an increase in ionic strength due to addition of monovalent salts such as NaCl.9 The kinetics of the surface fusion process have not been examined in great detail but are clearly accelerated by thermal and mechanical destabilization. For example, Kalb et al.9 observed lipid film formation within 6 min when a hydrophobic substrate was exposed to a 200 µM solution of palmitoyloleoylphosphatidylcholine (POPC) vesicles under conditions of steady laminar flow. Further exposure led to multilayer formation which required extensive rinsing to yield a monolayer. Others49 have shown that, at and above the phospholipid melting transition temperature, vesicle membranes are predisposed to defect formation, exposing the inner hydrophobic core and increasing the likelihood of fusion to a nearby hydrophobic surface. Of interest, Wenzl et al.10 noted that DPPC vesicles spontaneously fused at temperatures below Tm when less than 50 nm in diameter. They speculated (45) Stelze, M.; Weismu¨ller, G.; Sackmann, E. J. Phys. Chem. 1993, 97, 2974. (46) Singh, S.; Keller, D. J. Biophys. J. 1991, 60, 1401. (47) Johnson, M. E.; Berk, D. A.; Blankschtein, D.; Golan, D. E.; Jain, R. K.; Langer, R. S. Biophys. J. 1996, 71, 2656. (48) Evert, L. L.; Leckband, D.; Israelachvilli, J. N. Langmuir 1994, 10, 303. (49) Raudino, A.; Zucarello, F.; LaRosa, C.; Buenni, G. J. Phys. Chem. 1990, 94, 4217.
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Figure 6. AFM in contact mode performed under PBS and at room temperature on a series of surfaces composed of DPPC (A and B) and DLPC (C and D). Images were acquired in an immersed contact regime, using the constant height mode, with a horizontal resolution set at 512 pixels × 512 pixels and a scanning line speed of 4 Hz. The probe force was maintained at 0.5-1 nN throughout the imaging experiment. Average hole depths (arrow to arrow vertical distance) were 20 and 13 Å for DPPC (B) and DLPC (D) monolayers, respectively.
that fusion was facilitated due to the high internal stresses associated with small vesicle size. In our system, a temperature above the Tm of the phospholipid was selected to facilitate the fusion process. For DPPC, with a Tm of 41 °C, alkylated substrates were incubated with vesicles at 50 °C. In all other systems fusion was performed at room temperature given the Tm of 23 °C and -1 °C for DMPC and DLPC, respectively. While the appropriate choice of only a few parameters easily enhances vesicle fusion with a hydrophobic surface, a recognized disadvantage of this approach is an inability to predict a priori an optimal incubation period to minimize multilayer formation. Characteristically, after an arbitrary fusion period, copious surface rinsing is performed. The necessary volume of buffer required to mediate the removal of nonadsorbed liposomes has varied widely among reported procedures with values ranging from 10 to 100 fusion volumes.9,18,50,51 Excess rinsing raises the potential for damage to the supported monolayer, if fluid forces during buffer exchange are significant. We have observed that (50) Reinl, H. M.; Bayerl, T. M. Biochemistry 1994, 33, 14091. (51) Brink, G.; Schmitt, L.; Tampe´, R.; Sackmann, E. Biochim. Biophys. Acta 1994, 1196, 227.
excess lipids are effectively removed after exchange of approximately 24 fusion volumes, with suprisingly little subsequent degradation of the monolayer indicative of robust interlayer hydrophobic coupling. Radiotitration techniques have been used infrequently in surface analysis. Labeled DPPC proved useful in monitoring the effectiveness of the rinse procedure. However, the major significance of utilizing doped vesicles lay in providing independent confirmation of monolayer formation with molecular packing areas consistent with that reported for vesicle-based membrane systems at our fusion temperature of 50 °C. AFM imaging of supported membrane systems including membrane fragments,52,53 Langmuir-Blodgett films,52,54 and collapsed vesicles46 has been performed successfully by several groups. Most of these studies were conducted in air and in the case of DPPC bilayers (52) Meyer, E.; Howald, L.; Overney, R. M.; Heinzelmann, H.; Frommer, J.; Guntherodt, H. J.; Wagner, T.; Schier, H.; Roth, S. Nature 1991, 349, 398. (53) Worcester, D. L.; Miller, R. G.; Brynat, P. J. J. Microsc. (Oxford) 1988, 152, 817. (54) Zasadzinski, J. A.; Helm, C. A.; Longo, M. L.; Weisenhorn, A. L.; Gould, S. A.; Hansma, P. K. Biophys. J. 1991, 59, 755.
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mol %. Characteristically, phospholipids seldom exhibit ideal mixing behavior within self-assembled constructs.55 For example, lipids with identical head-group structure but exhibiting small differences in alkyl chain length will demonstrate significant phase separation in a binary mixture.48,56 Prior studies have demonstrated that the addition of cholesterol decreases the enthalpy of melting in direct proportion to cholesterol content, disappearing entirely at 50 mol % cholesterol.56 In this fashion, cholesterol acts as a plasticizer, increasing both membrane fluidity57 and stability.58 In support of these findings, holes were not detected in this study in either supported DPPC/Chol or DLPC/Chol monolayers. We suspect that the presence of cholesterol prevented monolayer contraction created by lateral film tension upon postfusion cooling. Another notable feature of cholesterol is its propensity to induce the spontaneous fusion of phospholipid bilayers.59 While this attribute may facilitate the disruption of the vesicle on contact with an alkylated surface with subsequent formation of a supported lipid film, it probably also increases the likelihood of liposome surface contamination. This was noted in our studies. The dimensions of surface mounds, as measured by AFM, were consistent with partially or hemifused liposomes in which only the outer leaflet of the liposome is fused to the SAM while the inner leaflet remains intact. Hemifusion of liposomes on planar phospholipid bilayers60 and on supports made hydrophobic by the adsorption of arachidonic acid61 has also been noted by other investigators. Our observation of membrane fusion with alkylsilane monolayers is of particular interest because, under similar experimental conditions, the fusion process appeared to be enhanced when cholesterol was added to the lipid vesicle. Chernomordik and colleagues62 have emphasized that membrane fusion in biological processes may involve the formation of specialized intermediates known as stalks and pores. In this model, the rate of fusion depends on the propensity of the corresponding monolayers of membranes to bend in the required direction. While proteins and peptides can control the bending energy of membrane monolayers, our data support the notion that that lipid composition actively contributes to the bending energy of these intermediates and is not a passive participant in the fusion process. Conclusions
Figure 7. Cholesterol-containing lipid vesicles fused to OTS glass and imaged by AFM under PBS and at room temperature. Analysis of supported DPPC/Chol (A), DMPC/Chol (B), and DLPC/Chol (C) films all revealed the presence of 150-200 nm surface “bumps” with an approximate height of 60 nm, consistent with fused liposomes. In part A, an adsorbed liposome (200 nm) is observed (white arrow). The horizontal (black arrows) and vertical lines are imaging artifacts.
demonstrated a thickness of approximately 60 Å.46 In our system, examination of a lipid monolayer in an aqueous environment reveals a measured leaflet thickness consistent with these investigations, as well as with other reported methods used to characterize the thickness of a DPPC bilayer. To our knowledge, AFM imaging of cholesterol-containing supported monolayers has not been reported. Cholesterol is present in cell membranes in levels up to 40
Using a combination of radiochemical titration techniques and AFM imaging in an aqueous environment, we have confirmed the formation of monolayer phospholipid membranes on smooth alkylated glass substrates. The composition and structural features of the lipid monolayer, as indicated by the molecular surface area and by the relative change in film thickness among an homologous lipid series, confirm the view that a membrane-mimetic lipid film can be produced on an alkylsilane self-assembled monolayer by vesicle fusion. (55) Shin, K.; Maeda, H.; Fujiwara, T.; Akutsu, H. Biochim. Biophys. Acta 1995, 1238, 42. (56) New, R. R. C. Liposomes: A Practical Approach; Oxford University Press: New York, 1992. (57) Rubenstein, J. L.; Smith, B.; McConnell, H. M. PNAS (USA) 1979, 76, 15. (58) Bedu-Addo, F. K.; Tang, P.; Xu, Y.; Huang, L. Pharm. Res. 1996, 13, 718. (59) Vogel, S. S.; Chernomordik, L. V.; Zimmerberg, J. J. Biol. Chem. 1992, 267, 25640. (60) Perin, M. S.; MacDonald, R. C. J. Membr. Biol. 1989, 109, 221. (61) Vikholm, I.; Peltonen, J.; Teleman, O. Biochim. Biophys. Acta 1995, 1233, 111. (62) Chermnomordik, L.; Kozlov, M. M.; Zimmerberg, J. J. Membr. Biol. 1995, 146, 1.
Supported Phospholipid Monolayers
Acknowledgment. This work was supported by the Whitaker Foundation and the NIH (Grant HL56819). E.L.C. is a Clinician-Scientist of the American Heart Association. The authors wish to acknowledge Professor Joseph A. Gardella, Jr., and Dr. Richard Nowak at SUNY Buffalo for ESCA measurements and Professor Lawrence
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A. Bottomley at the Georgia Institute of Technology, Department of Chemistry, for assistance with AFM imaging. LA980077P
