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Influence of Surface Properties of Mixed Monolayers on Lipolytic Hydrolysis G. H. Peters,* U. Dahmen-Levison,† K. de Meijere,† G. Brezesinski,† S. Toxvaerd,‡ H. Mo¨hwald,† A. Svendsen,§ and P. K. J. Kinnunen| Department of Chemistry, Membrane and Statistical Physics Group (MEMPHYS), Technical University of Denmark, Building 206, DK-2800 Lyngby, Denmark, Max Planck Institute of Colloids and Interfaces, Am Mu¨ hlenberg 2, D-14476 Golm/Potsdam, Germany, Chemistry Department III, H. C. Ørsted Institutet, University of Copenhagen, Universitetsparken 5, DK-2100 Copenhagen Ø, Denmark, Novo Nordisk A/S, Novo Alle´ 1, DK-2880 Bagsvaerd, Denmark, and Department of Medical Chemistry and Department of Physiology, Institute of Biomedicine, University of Helsinki, Helsinki, Finland Received May 28, 1999. In Final Form: November 10, 1999 Fluorescence microscopy, surface potential, and activity measurements were used to investigate the influence of fatty acids and fatty alcohols on the lipolytic activity of several lipases. We have determined the lateral lipid distribution and interfacial properties of Langmuir mixed monolayers composed of 1,2didecanoylglycerol/eicosanoic acid or 1,2-didecanoylglycerol/1-octadecanol molecules and have measured lipase activities toward these films. Enzymatic activities are remarkably influenced by the addition of fatty acid. Activity decreases continuously up to a mole fraction of ≈ 0.1 fatty acid, where phase separation and a change in surface potential are observed. Higher concentrations of fatty acid have only marginal effects on the lipase activities. The relative activity between the different lipases varies substantially, and there is an indication that the level of inhibition correlates with the isoelectric point (pI) of the enzymes. A simpler mechanism is observed by the addition of fatty alcohol. Within the concentration range studied, 1-octadecanol is immiscible in the diacylglyceride matrix, forming liquid-condensed domains. The inhibitory effect is related to the reduction of available diacylglyceride area to the enzyme. Direct imaging of the lipolytic hydrolysis of these monolayers show that relatively small domains are formed, suggesting that the enzyme preferentially acts on pure diacylglyceride patches.
Introduction Lipases (acylglycerol acylhydrolase, EC 3.1.1.3) constitute a ubiquitous class of enzymes, which are recognized as extremely versatile enzymes. Their capability for catalyzing a wide variety of reactions, which include transesterification, stereospecific synthesis of compounds, and so forth, allows for a wide range of industrial application. The activity of these lipolytic enzymes increases substantially when adsorbed onto a lipid-water interface. The overall catalytic hydrolysis involves adsorption of the enzyme from the bulk aqueous phase to the surface, followed by catalytic action at the interfacial plane.1,2 Theories have been proposed that either emphasize the structure of the interfacial lipid-water region (“substrate theory”) or conformational changes in the enzyme upon adsorption to the interface (“enzyme theory”).3-8 Several lipase structures have been resolved * To whom correspondence should be addressed at the Technical University of Denmark. Tel.: (+45) 4525 2385. Fax: (+45) 4593 4808. E-mail:
[email protected]. † Max-Planck Institute of Colloids and Interfaces. ‡ University of Copenhagen. § Novo Nordisk A/S. | University of Helsinki. (1) Derewenda, U.; Swenson, L.; Wei, Y.; Green, R.; Kobos, P. M.; Joerger, R.; Haas, M. J.; Derewenda, Z. S. J. Lipid Res. 1994, 35, 524. (2) Brzozowski, A. M.; Derewenda, U.; Derewenda, Z. S.; Dodson, G. G.; Lawson, D. M.; Turkenburg, J. P.; Bjorkling, F.; Huge-Jensen, B.; Patkar, S. A.; Thim, L. Nature 1991, 351, 491. (3) Van Tilbeurgh, H.; Egloff, M. P.; Martinez, C.; Rugani, N.; Verger, R.; Cambillau, C. Nature 1991, 351, 814. (4) Muderhwa, J. M.; Brockman, H. L. J. Biol. Chem. 1992a, 267, 24184. (5) Thuren, T. FEBS Lett. 1988, 229, 95. (6) Momsen, W. E.; Brockman, H. L. J. Biol. Chem. 1981, 256, 6913. (7) Wells, X. Biochemistry 1974, 11, 1030.
crystallographically, revealing that the active site is shielded from the solvent by an R-helical loop (“lid”), which, upon activation, rolls over to allow access of the substrate to the active site. In the inactive conformation of the enzyme, the exposed hydrophobic surface is minimal, but, as the lid rolls over, a large hydrophobic patch is exposed that interacts favorably with the nonpolar interface.9,10 It has been shown that the charge distribution close to the active site is important in stabilizing the active conformation of several lipases,11-13 which may also affect the selectivity toward substrates and the sensitivity to inhibitors. Inhibition of the lipolytic hydrolysis by triacylglycerol lipases occurs in the presence of micellular concentrations of bile salts,14-16 fatty acids, fatty alcohols, halogenated phenylamines, or hydrocarbons.17-19 Mattson et al.18 have performed a detailed study on the inhibition of lipolysis by normal alcohols. The authors concluded that the (8) Wells, X. Biochemistry 1974, 13, 2248. (9) Derewenda, Z. S. Nat. Struct. Biol. 1995, 2, 347. (10) Dodson, G. G.; Lawson, D. M.; Winkler, F. K. Faraday Discuss. 1992, 93, 95. (11) Peters, G. H.; Olsen, O. H.; Svendsen, A.; Wade, R. Biophys. J. 1996, 71, 119. (12) Peters, G. H.; Toxvaerd, S.; Olsen, O. H.; Svendsen, A. Protein Eng. 1997, 10, 137. (13) Holmquist, M.; Martinelle, M.; Berglund, P.; Clausen, I. G.; Patkar, S.; Svendsen, A.; Hult, K. J. Prot. Chem. 1993, 12, 749. (14) Pie´roni, G.; Gargouri, Y.; Sarda, L.; Verger, R. Adv. Colloid Interface Sci. 1990, 32, 341. (15) Verger, R.; Pattus, F.; Pie´roni, G.; Riviere, C.; Ferrato, F.; Leonardi, J.; Dargent, B. Colloids Surf. 1984, 10, 163. (16) Borgstro¨m, B. Biochim. Biophys. Acta 1977, 488, 381. (17) Ferreira, G. C.; Patton, J. S. J. Lipid Res. 1990, 31, 889. (18) Mattson, F. H.; Volpenheim, R. A.; Benjamin, L. J. Biol. Chem. 1970, 245, 5335. (19) Comai, K.; Sullivan, A. C. J. Pharm. Sci. 1982, 71, 418.
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inhibition of short-chain alcohols is caused by interfacial adsorption rather than the formation of an enzymealcohol complex.18 Some putative inhibitors exhibit an activation at low concentrations and inhibition at high concentrations.20 For instance, alcohols and detergents can inhibit the catalyzed hydrolysis of phospholipase A2 at high concentrations, whereas at low concentrations, they can stimulate the enzyme.21 The inhibitory action may involve adsorption at the surface or incorporation into the substrate matrix17,21 and is related to the amphiphilic nature of the additives, which may act as spacers to facilitate the adsorption of the enzyme at the lipid/water interface.20 Effects of fatty acids on both interfacial adsorption and subsequent catalysis have been observed for pancreatic carboxylester lipase.22 These inhibitory effects indicate that an important factor in the regulation of lipolytic enzymes in vitro and in vivo is the lateral distribution and packing of lipid species in the interfacial plane6,23,24. An understanding of the regulation of lipolytic hydrolysis requires that the binding of lipases onto the lipid interface and the catalytic reaction each be investigated with respect to the chemical composition and physical structure of the substrate. A useful approach has been to eliminate the bulk lipid phase and to control surface area by using monomolecular films (Langmuir monolayers) of insoluble lipids at the air-water interface.14,15 Model studies using Langmuir films are beginning to reveal the relationship between enzyme kinetics and lipid properties such as interfacial properties, surface composition, and concentration.25,26 Structural ordering in monolayers is generally determined from synchrotron X-ray scattering experiments27-31 or imaging techniques such as Brewster angle microscopy, fluorescence microscopy, and transmission electron microscopy.32-39 These microscopic techniques have been widely utilized in areas such as membranes and monolayer textures. Using Brewster angle or fluorescence microscopy, the direct observation of phase equilibrium between liquid expanded and condensed phases is now common practice. Recently, fluorescence microscopy has been introduced to study the interaction of phospholipase A2 with monolayers (20) Jain, M. K.; Berg, O. G. Biochim. Biophys. Acta 1989, 1002, 127. (21) Ransac, S.; Rivie´re, C.; Soulie´, J. M.; Gancet, C.; Verger, R.; de Haas, G. H. Biochim. Biophys. Acta 1990, 1043, 57. (22) Muderhwa, J. M.; Schmid, P. C.; Brockman, H. L. Biochemistry 1992, 31, 141. (23) Smaby, J. M.; Muderhwa, J. M.; Brockman, H. L. Biochemistry 1994, 33, 1915. (24) Thuren, T.; Wilcox, R. W.; Sisson, P.; Waite, M. J. Biol. Chem. 1991, 266, 4853. (25) Rubingh, D. N. Curr. Opin. Colloid Interface Sci. 1996, 1, 598. (26) Turro, N. J.; Xue-Gong, L.; Ananthapadmanbhan, K. L.; Aronson, M. Langmuir 1995, 11, 2525. (27) Lin, B.; Smith, M. C.; Bohanon, T. M.; Ice, G. E.; Dutta, P. Phys. Rev. Lett. 1990, 65, 191. (28) Kenn, R. M.; Bo¨hm, C.; Bibo, A. M.; Peterson, I. R.; Mo¨hwald, H.; Als-Nielsen, J.; Kjaer, K. J. Phys. Chem. 1991, 95, 2092. (29) Schwartz, D. K.; Schlossman, M. I.; Pershan, P. S. J. Chem. Phys. 1992, 96, 2356. (30) Li, M.; Acero, A. A.; Huang, Z.; Rice, S. A. Nature 1994, 367, 151. (31) Weinbach, S. P.; Kjaer, K.; Bouwman, W. G.; Gru¨bel, G.; Legrand, J. F.; Als-Nielsen, J.; Lahav, M.; Leiserowitz, L. Science 1994, 264, 1566. (32) Knobler, C. M. Adv. Chem. Phys. 1990, 397. (33) Mo¨hwald, H. Rep. Prog. Phys. 1993, 56, 653. (34) Patino, J. M. R.; Sanchez, C. C.; Nino, M. R. R. Langmuir 1999, 15, 2484. (35) Mazur, A. W.; Burns, J. L.; Hiler, G. D., II; Spontak, R. J. J. Phys. Chem. 1993, 97, 11344. (36) Ruiz-Garcia, J.; Qui, T.; Tsao, M. W.; Marshall, G.; Knobler, C. M.; Overbeck, G. A.; Mo¨bius, D. J. J. Phys. Chem. 1993, 97, 6955. (37) Schwartz, D. K.; Knobler, C. M. J. Phys. Chem. 1993, 97, 8849. (38) Overbeck, G. A.; Mo¨bius, D. J. J. Phys. Chem. 1993, 97, 7999. (39) Riviere, S.; Henon, S.; Meunier, D.; Schwartz, D. K.; Tsao, M. W.; Knobler, C. M. J. Chem. Phys. 1994, 101, 10045.
Peters et al.
of DPPC, DMPC, and DPPE40,41 and with mixed monolayers of phosphatidylcholine, lecithin, and fatty acid.42 Visualization of lipid phase separation as well as enzymatic hydrolysis can provide valuable information concerning the effects of lipid compositions, metal ions, pH, and so forth, on enzyme activity. A similar approach is used in the present study to investigate the catalytic hydrolysis and to elucidate the nature of the inhibition of triacylglycerol lipases by fatty acids and fatty alcohols. Experimental Section Materials. All reagents and solvents were of the highest purity available. Milli-Q water (18 M ohm‚ cm) was used for all aqueous solutions and dilutions. Chloroform used as a spreading solvent was analytical grade and a product of Merck, Darmstadt, FRG. 1,2-Didecanoyl-rac-glycerol, C23H44O5, (D-6389; Lot 34H8488; ≈ 99% pure), 1-octadecanol, C18H38O1, (S-5751; Lot 24H1025; ≈ 99% pure), NBD, C36H62N5O11P1, (L-R-phosphatidylcholine-β-(NBDaminohexanoyl)-γ -palmitoyl; P-3412; Lot 108F8459) and Tris (Tris-[hydroxymethyl]-aminomethane; T-1503; Lot 122H5608) were purchased from Sigma, St. Louis, MO. Eicosanoic acid, C20H40O2, (puriss. grade) was a product of Fluka, Buchs, Switzerland and was used as supplied. The following lipases were purchased from Sigma, St. Louis, MO.: Candida Rugosa (900 units/mg solid; L-1754; Lot 100H0482), Rhizomucor Miehei (1000 units/mg solid), and Pseudomonas Cepacia (12230 units/ml). Rhizopus Delemar lipase was ordered from Seikagaku Kogyo, Tokyo, Japan (600 units/mg solid; Lot 17001). Activity Measurements. Lipase activities were determined in a zero-order Teflon trough14 using the following enzyme concentrations: 1 unit of Rhizomucor Miehei lipase, 1 unit of Rhizopus Delemar lipase, 50 units of Candida Rugosa lipase, and 50 units of Pseudomonas Cepacia lipase. The trough consisted of reservoir and reaction compartments, which were connected by a small channel (4-mm width). The dimensions of the reservoir and reaction chambers were (395 × 150 × 20 mm) and (70 × 150 × 20 mm), respectively. The reaction compartment was equipped with two magnetic stirrers. Didecanoylglycerol and eicosanoic acid or didecanoylglycerol and 1-octadecanol were mixed in chloroform in the indicated molar ratios and were subsequently spread from this solution onto the subphase at a temperature of 25 °C. The subphase was 10 mM Tris buffer containing 0.1mM EDTA (ethylenediaminetetraacetic acid). The addition of EDTA was performed to remove any traces of Ca2+ and to avoid any interference by calcium ions. Lipase activity is influenced by Ca2+ ions,43 which could be due to the interaction with the enzyme (although there is no Ca2+ binding site in the protein) or the lipid interface. As a divalent ion, it is expected that the calcium ion interacts with the fatty acid molecules in the monolayer (as a counterion in the double layer). The buffer was prepared by dissolution of salts in 10-times concentration, adjusting the pH to 8.0 by adding 1 N HCl solution and filtering through a 22-µm filter. Prior to each experiment, the solution was diluted with milli-Q water to the proper concentration for the monolayer subphase. Surface pressure measurements were performed with a KSV5000-3 (40) Grainger, D. W.; Reichert, A.; Ringsdorf, H.; Salesse, C. FEBS Lett. 1989, 252, 73. (41) Grainger, D. W.; Reichert, A.; Ringsdorf, H.; Salesse, C. Biochim. Biophys. Acta 1990, 1023, 365. (42) Reichert, A.; Ringsdorf, H.; Wagenknecht, A. Biochim. Biophys. Acta 1992, 1106, 178. (43) Duinhoven, S.; Poort, R.; van der Voet, G.; Agterof, W. G. M.; Norde, W.; Lyklema, J. J. Colloid Interface Sci. 1995, 170, 351.
Mechanism of Inhibition of Lipase Catalysis
manufactured by KSV Instruments, Helsinki. The experiments were conducted as follows: Lipid solution was spread on the subphase to reach an initial surface pressure close to the desired value of ≈ 25 mN/m. Small deviations from the actual value were compensated by compressing the film. The monolayer was maintained at this pressure for 30-60 min to reach thermodynamic equilibrium. Activity measurements were started by injecting the lipase into the subphase of the reaction compartment (1-2 mm below the surface). Enzymatic hydrolysis of the didecanoylglycerol molecules yields glycerol and fatty acid. These short-chain molecules cannot form a stable monolayer and are water soluble. The additives, eicosanoic acid or 1-octadecanol, have longer hydrocarbon chains and hence form stable monolayers. This approach allows the study of lipase activity in the presence of a fatty acid or fatty alcohol. To maintain constant surface pressure, the barrier automatically compresses the film. The speed by which the barrier moves is a measure of the activity. For all lipases, an initial lag time was observed. For a pure didecanoyl-rac-glycerol monolayer, the barrier speed initially increased and, depending on the lipase, a constant barrier speed was observed after approximately 10 min. Each profile was fitted to a hyperbolic function to determine the maximum (constant) speeds, which were as follows: Candida Rugosa lipase, 0.5 mm/s; Pseudomonas Cepacia lipase, 0.6 mm/s; Rhizomucor Miehei lipase, 0.2 mm/s; and Rhizopus Delemar lipase, 0.2 mm/s. Based on these barrier speeds, the hydrolysis rates are Candida Rugosa lipase, 6.1 pmol FFA (free fatty acid)/cm2/sec; Pseudomonas Cepacia lipase, 7.3 pmol FFA/cm2/sec; Rhizomucor Miehei lipase, 2.4 pmol FFA/cm2/sec; and Rhizopus Delemar lipase, 2.4 pmol FFA/cm2/sec. All reported activity data are normalized by the barrier speed measured for a pure diacylglyceride film. Fluorescence Microscopy. Images were generated from the light emitted by the NBD fluorescent probe, which was added to the spreading solution in small quantities (0.5-1.0 mole percent). The NBD molecules preferentially partition into the fluid phase, causing a bright image in the microscope. As the phase separation occurs and solidlike domains are formed, NBD molecules are pushed out of the solid phase, and the domains appear as a dark region in the image. Phase equilibria of didecanoylglycerol/ eicosanoic acid and didecanoylglycerol/octadecanol monolayers were monitored using a 250 × 60 × 6 mm firstorder Teflon trough (R&K, Germany), applying similar spreading conditions as described above. The trough used in the experiment was smaller than the trough used in the activity measurements. This was required by the experimental setup to allow the continuous observation of the film. Hydrolysis was monitored by spreading lipid solution onto the subphase (10mM Tris buffer with 0.1mM EDTA; pH ) 8) containing the lipase and continuously viewing the monolayer through the microscope. Since the enzymatic reaction started during spreading, it was difficult to determine exactly the initial surface pressure. However, only small amounts of lipid solution were required (≈ 30 µL) to reach surface pressures close to the desired value of 25 mN/m. Monolayers were viewed from above through a microscope (Zeiss Axiotron), which was equipped with several objectives (LD 10×/0.3, LD 20×/ 0.4, LD 50×/0.5). The light source was a Hg lamp (HBO 50 W), where the excitation beam was obtained through a filter transmitting in the range of 450-490 nm. The image was collected by a SIT camera (Hamamatsu C2400) and viewed on the display screen. During compression, the images were stored by a video recorder. Some images
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were digitized and enhanced on a personal computer using the NIH 1.42 Image software. Surface Potential. Surface potentials were measured using the ionizing electrode method.44 The ionizing source was americium-241 (Amersham International plc, Amersham Place, Little Chalfont, Bucks HP7 9NA, U.K.), and the reference electrode was a 2.7 kΩ KCl electrode; FLEX-REF (World Precision Instr., Inc., 175 Sarasota Center Boulevard, Sarasota FL 34240-9258, U.S.A.). To compare to the results of the activity measurements, the same conditions were chosen; that is, the same subphase and the same composition of the mixed monolayers. Potential data of didecanoylglycerol/eicosanoic acid and didecanoylglycerol/octadecanol monolayers as a function of surface pressure were determined using a zero-order Teflon trough.14 Reservoir and reaction compartments having areas of 40 314 and 12 660 mm2, respectively, were connected by a small channel (4-mm width). Preparation of the lipid solutions and spreading conditions were similar to those described above. Solutions were spread onto the subphase (10 mM Tris buffer with 0.1 mM EDTA; pH ) 8) at a temperature of 25 °C, resulting in an initial pressure of
