Langmuir 1999, 15, 1731-1737
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Biomembrane Mimetic Surfaces by Phospholipid Self-Assembled Monolayers on Silica Substrates Zhihao Yang† and Hyuk Yu* Department of Chemistry, University of Wisconsin, Madison, Wisconsin 53706 Received July 8, 1998. In Final Form: December 3, 1998 A method of covalently bonding phospholipid molecules to silica substrates followed by loading with free phospholipids is demonstrated to form well-organized and stable phospholipid self-assembled monolayers (SAMs). Surfaces of such SAMs, which structurally mimic the aqueous sides of phospholipid bilayer membranes, are characterized with X-ray photoelectron spectroscopy and atomic force microscopy. Dynamics of phospholipids and an adsorbed protein, lipase, in the SAMs are probed with the technique of fluorescence recovery after photobleaching (FRAP), in terms of lateral diffusion of both phospholipids and protein molecules. The lateral diffusion coefficient of the free lipids in the SAMs is found as (1.9 ( 0.4) × 10-9 cm2/s whereas that of lipase is (2.7 ( 0.4) × 10-10 cm2/s; both are within an order of magnitude of those in cell membranes. The esterase activity of lipase on the SAM surfaces is confirmed by the hydrolysis reaction of a substrate, umbelliferone stearate.
* To whom correspondence should be addressed. Phone: (608) 262-3082. Facsimile: (608) 262-9918. E-mail:
[email protected]. † Current address: Eastman Kodak Company, 1999 Lake Ave. B82A, Rochester, NY 14650-2121.
Venkataram8 to bond covalently phospholipids on a silica gel surface for chromatography applications, has greatly inspired our interest in further developing a stable and robust self-assembled lipid monolayer on solid surfaces to mimic the functionality of biomembranes, especially the surface activation of membrane-bound enzymes by the lipid self-assembled monolayers. One of the most distinctive and intriguing features of lipolytic enzymes is their activation by such interfaces as cell membranes.9 Lipase, a lipolytic enzyme with esterase activity, is capable of rapid hydrolysis of sparingly soluble substrates, such as fats and oils, in an amphiphilic environment. In recent years, lipases have become a very important class of enzyme used in asymmetric organic synthesis because of their special stereoselectivity.10,11 On the other hand, lipases are found to be insensitive toward soluble esters in solution, and only when sparing soluble esters exceed their critical micelle concentration, do lipases exhibit a dramatic increase in their enzymatic activity. Such an interfacial activation phenomenon of lipases was first recognized by Holwerda et al.12 More recently, by determining lipase structures, Lawson and co-workers attributed the interfacial activation of lipase to a conformational change of the enzyme to expose its active site to the surface once it encounters a lipid micelle.13 Thus, a biomembrane mimetic environment, such as a lipid/ water interface, is essential for lipase to be enzymatically active. In this study, we focus on a method to form a selfassembled monolayer (SAM) of phospholipid molecules, well organized and stable, by covalently bonding a certain fraction of them to a silica substrate followed by loading with free lipids, which then self-assemble laterally with
(1) Sackmann, E. Science 1996, 271, 43. (2) Puu, G.; Gustafson, I.; Artursson, E.; Ohlsson, P.-A° . Biosens. Bioelectron. 1995, 10, 463. (3) Cai, S.; McAndrew, R. S.; Leonard, B. P.; Chapman K.; Pidgeon, C. J. Chromatogr. A 1995, 696, 49. (4) Pidgeon, C.; Ong, S.; Liu, H.; Qiu, X.; Pidgeon, M.; Dantzig, A. H.; Munroe, J.; Hornback, W. J.; Kasher, J. S.; Glunz, L.; Szczerba, T. J. Med. Chem. 1995, 38, 590. (5) Tamm, L. K.; Bo¨hm, C.; Yang, J.; Shao, Z.; Hwang, J.; Edidin, M.; Betzig, E. Thin Solid Films 1996, 284, 813. (6) Stelzle, M.; Weissmu¨ller, G.; Sackmann, E. J. Phys. Chem. 1993, 97, 2974. (7) Heckl, W. M.; Ho¨rber, H. J. K.; Binnig, G. Langmuir 1989, 5, 1433.
(8) Pidgeon, C.; Venkataram, U. V. Anal. Biochem. 1989, 176, 36. (9) Verger, R. Methods Enzymol. 1980, 64, 340. (10) Rogalska, E.; Ransac, S.; Verger, R. J. Biol. Chem. 1990, 265, 20271. Rogalska, E.; Ransac, S.; Verger, R. J. Biol. Chem. 1992, 268, 792. Rogalska, E.; Nury, S.; Douchet, I.; Verger, R. Chirality 1995, 7, 505. (11) Kazlauskas, R. J.; Weissfloch, A. N. E.; Rappaport, A. T.; Cuccia, L. A. J. Org. Chem. 1991, 56, 2656. (12) Holwerda, K.; Verkade, P. E.; de Willingen, A. H. A. Recl. Trav. Chim. Pays-Bas 1936, 55, 43. (13) Lawson, D. M.; Brzozowski, A. M.; Dodson, G. G. Curr. Biol. 1992, 2, 473.
Introduction Cell membranes interacting with all types of biomolecules are responsible for many important biological processes, including molecular recognition, cell adhesion, intercellular communication, and enzymatic catalysis. For both scientific importance and practical applications, the supported membranes (lipid bilayers and monolayers) on solids are of great interest, since they enable biofunctionalization of inorganic solids and polymeric materials, and provide a natural environment for the interacting biomolecules retaining their functionality.1,2 The supported lipid-protein membranes are also of importance in many applications, including biomolecule separation,3 drug delivery,4 design of biosensors,5,6 and biomimetic catalysis. The most commonly used methods of constructing membrane assemblies on surfaces are monolayer transfer,1 that is, the Langmuir-Blodgett technique, and vesicle spreading. There are two major limitations for the transferred monolayer systems: (1) Thermodynamic instability renders them suffering from lack of long-term durability; (2) the transferring methods require the supporting substrates to be planar, which will not provide large enough surface area for many important applications, such as enzymatic catalysis. Thus, direct graft of phospholipids to solid substrate to form a highly packed monolayer holds great promise in biomembrane mimetic applications.7 A method, first reported by Pidgeon and
10.1021/la980839g CCC: $18.00 © 1999 American Chemical Society Published on Web 02/11/1999
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the bonded ones as the template. It is expected that such a monolayer resembles the outer half-plane of phospholipid bilayer membranes not only structurally but also dynamically in terms of intralamellar lateral diffusion of monolayer components. By the outer half-plane we mean that the polar heads of lipids are directed toward the aqueous medium while the hydrophobic tails are linked or directed to the substrate surface. The enzymatic viability is to be examined by an ester hydrolysis reaction with the aid of lipase from Pseudomonas cepacia. Experimental Section Materials. Monomyristoyl lysolecithin (lyso-PC) and 1-myristoyl-2-12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoylsn-glycero-3-phosphoethanolamine (NBD-PE) in a CHCl3 solution were purchased from Avanti Polar Lipids, Inc. (Alabaster, Alabama), and dilauroylphosphatidylcholine (DLPC) was from Sigma. Pseudomonas cepacia lipase (Amano LPL-200S, molecular weight of 33 089 Da) was a product from Amano Pharmacetical Co. (Nagoya, Japan.) Three (3-aminopropyl)silane (APS) derivatives, (3-aminopropyl)triethoxylsilane (APTES), (3-aminopropyl)diethoxylmethylsilane (APDEMS), and (3-aminopropyl)ethoxyldimethylsilane (APEDMS), were purchase from Gelest, Inc. (Tullytown, PA). Dicyclohexylcarbodiimide (DCC), 1,12-dodecanedicarboxylic acid, 4-N,N-dimethylaminopyridine (DMAP), carbonyldiimidazole (CDI), fluorescein isothiocyanate (FITC), and all anhydrous solvents were purchased from Aldrich Chemical Co. The DMAP was recrystallized from diethyl ether before use. Nucleosil-300-7NH2 was a product from Macherey-Nagel (Germany). The water used throughout the experiment was purified with a Milli-Q system from Millipore Co. with its initial resistivity better than 17 MΩ/cm. Synthesis of 1-Myristoyl-2-ω-carboxylmyristoyl-sn-3glycerophosphocholine (DMPC-COOH). The DMPC-COOH was synthesized by following the procedure of Pidgeon and Vankataram8 with minor modifications. Briefly, 10.0 g (0.038 mol) of 1,12-dodecanedicarboxylic acid was dissolved in 200 mL of anhydrous tetrahydrofaran (THF) in a dry 500 mL roundbottom flask at 40 °C, and 8.1 g (39 mmol) of DCC was dissolved in 20 mL of anhydrous THF in a dry beaker. The DCC solution was added to the reaction flask dropwise with a pipet while being purged with dry nitrogen, and a white suspension resulted. The reaction flask was then sealed under nitrogen and kept stirring at room temperature for 15 h. Then, 500 mL of acetone was added and filtered through a sintered glass funnel. The filtrate was slightly cloudy and stored in a refrigerator overnight. The cloudy solution was filtered again, and the solid product was collected and dried under vacuum (
