7400
Langmuir 2008, 24, 7400-7407
Competition between Lipases and Monoglycerides at Interfaces Pedro Reis,†,‡ Krister Holmberg,*,‡ Reinhard Miller,§ Jurgen Kra¨gel,§ Dmitri O. Grigoriev,§ Martin E. Leser,† and Heribert J. Watzke† Nestle´ Research Center, CH-1000 Lausanne 26, Switzerland, Department of Chemical and Biological Engineering, Applied Surface Chemistry, Chalmers UniVersity of Technology, SE-412 96 Go¨teborg, Sweden, and Max-Planck-Institut fu¨r Kolloid- and Grenzfla¨chenforschung, Max-Planck-Campus, D-14476 Golm, Germany ReceiVed February 19, 2008. ReVised Manuscript ReceiVed April 23, 2008 Tensiometry (the pendant drop technique), interfacial shear rheology, and ellipsometry have been used to study the effect of polar lipids that are generated during fat digestion on the behavior of lipases at the oil-water interface. Both Sn-1,3 regiospecific and nonregiospecific lipases have been used, and a noncatalytically active protein, β-lacloglobulin, has been used as reference in the interfacial shear rheology experiments. The results from the pendant drop measurements and the interfacial rheology studies were in agreement with each other and demonstrated that the Sn-2 monoglyceride, which is one of the lipolysis products generated when a Sn-1,3 regiospecific lipase catalyzes triglyceride hydrolysis, is very interfacially active and efficiently expels the enzyme from the interface. Ellipsometry conducted at the liquid-liquid interface showed that the lipase forms a sublayer in the aqueous phase, just beneath the monoglyceride-covered interface. Sn-1/3 monoglycerides do not behave this way because they are rapidly degraded to fatty acid and glycerol and the fatty acid (or the fatty acid salt) does not have enough interfacial activity to expel the lipase from the interface. Since the lipases present in the gastrointestinal tract are highly Sn-1,3 regiospecific, we believe that the results obtained can be transferred to the in ViVo situation. The formation of stable and amphiphilic Sn-2 monoglycerides can be seen as a self-regulatory process for fat digestion.
Introduction Interfaces play an important role in biology. The majority of biological events occur at interfaces rather than in bulk. One very important interfacial process is fat digestion, which occurs not only in animals but also in plants and microorganisms.1–3 Lipases and phospholipases are the enzymes involved in fat digestion. The most widely studied lipases are water-soluble,4 but many of their natural substrates are insoluble in water. It is, therefore, well-established that in order for a lipase to attain high activity a lipid-water interface is required.5 It has been demonstrated that binding of the lipase to the interface is a crucial step in biocatalysis.6 Proteins in general adsorb strongly to interfaces, and although the process is thermodynamically reversible, the desorption is often so slow that the adsorption can be regarded to be an irreversible process.7 Pancreatic lipase, however, being a relatively small protein, has been shown not to be irreversibly adsorbed at the oil-water interface.8 It has been known for a long time that low molecular weight surfactants can displace proteins from interfaces.9 Lipase* To whom correspondence should be addressed. E-mail: kh@ chalmers.se. † Nestle´ Research Center. ‡ Chalmers University of Technology. § Max-Planck-Institut fu¨r Kolloid- and Grenzfla¨chenforschung. (1) Jaeger, K. E.; Ransac, S.; Dijkstra, B. W.; Colson, C.; van-Heuvel, M.; Misset, O. FEMS Microbiol. ReV. 1994, 15, 29. (2) Gilbert, E. J. Enzyme Microb. Technol. 1993, 15, 634. (3) Wohlfahrt, S.; Jaeger, K. E. BioEngineering 1993, 9, 39. (4) Stamatis, H.; Xenakis, A.; Kolisis, F. N. Biotechnol. AdV. 1999, 17, 293. (5) Van-Tilebeurgh, H.; Egloff, M. P.; Rugani, C. M.; Verger, R.; Cambillau, C. Nature 1993, 362, 814. (6) Cordle, R. A.; Lowe, M. E. J. Lipid Res. 1998, 39, 1759. (7) Fainerman, V. B.; Miller, R.; Ferri, J. K.; Watzke, H.; Leser, M. E.; Michel, M. AdV. Colloid Interface Sci. 2006, 123, 163. (8) Haiker, H.; Lengsfeld, H.; Hadvary, P.; Carriere, F. Biochim. Biophys. Acta 2004, 1682, 72. (9) Mackie, A. R. Curr. Opin. Colloid Interface Sci. 2004, 9, 357.
catalyzed hydrolysis of triglycerides yields diglycerides and fatty acids (or fatty acid salts). The diglycerides are further degraded to monoglycerides and fatty acids. Provided the enzyme is Sn1,3 regiospecific, which is the case for the normal gastric and pancreatic lipases, the monoglyceride formed is the ester of the secondary hydroxyl group of glycerol, and this species is relatively stable toward further degradation. We have recently shown that monoglycerides can be the most interfacially active molecules generated during lipolysis.10 We have also demonstrated that the rate of fat digestion can be considerably decreased in a model gastrointestinal system by enrichment of the test meal with an Sn-2 monoglyceride11 (provided that acyl migration does not occur). In the present work, we have used tensiometry, interfacial shear rheology, and ellipsometry to study the effect of the individual lipolytic products on the interfacial behavior of lipases with different regiospecificity (Sn-1,3 or nonregiospecific) and origin (Rhizomucor miehei, Candida rugosa, and pancreatic lipase). A model globular protein, β-lactoglobulin (BLG), is used as reference in order to be able to separate the effect related to competition for the interface from that related to influence on the biological activity due to interactions between the amphiphilic lipolysis products and the protein. BLG also has the capacity to bind free fatty acids. We believe that our results will lead to a better understanding of the biophysics of fat digestion with a long-term view toward new approaches to modulate the lipolysis rate.
Materials and Methods Chemicals. Decane (>95.0%), caprylic acid C2875 (C8, >99%), lipase from Rhizomucor miehei (4210 units/mg solid), lipase from porcine pancreas (100-400 units/mg solid), and lipase from Candida (10) Reis, P.; Miller, R.; Leser, M. E.; Watzke, H.; Fainerman, V. B.; Holmberg, K. Langmuir 2008, 24, 5781. (11) Reis, P.; Rabb, T.; Chuat, J. Y.; Leser, M. E.; Miller, R.; Watzke, H.; Holmberg, K. Food Biophys., submitted for publication, 2008.
10.1021/la800531y CCC: $40.75 2008 American Chemical Society Published on Web 06/12/2008
Lipase and Monoglyceride Competition at Interfaces
Langmuir, Vol. 24, No. 14, 2008 7401
rugosa (10 699 units/ mg solid) were obtained from Sigma, St. Louis, MO. The lipase preparations from Rhizomucor miehei and Candida rugosa are rather pure but contain traces of other compounds (propanediol, potassium sorbate, sodium benzoate, sorbitol) that are used to stabilize the lipophylized form of the enzyme. The lipase from porcine pancreas is a crude preparation that contains other substances present in the duodenum (bile, proteases, colipase, etc). Sn-1/3 monocaprylin (Sn-1/3 MC8, 99%), Sn-2 monocaprylin (Sn-2 MC8, 99%), dicaprylin (DC8, 99%), Sn-1/3 monopalmitin (Sn-1/3 MC16, 99%), and Sn-2 monopalmitin (Sn-2 MC16, 99%) were obtained from Larodan, Malmo¨, Sweden. BioPure β-lactoglobulin (BLG) was supplied by Davisco, Le Sueur, MN. Pendant Drop Technique. Interfacial tension and viscoelasticity were recorded on a tensiometer from Teclis/IT Concept (Tracker). A syringe of 250 µL from Hamilton 1001 TLLWS with a needle of 1.83 mm diameter was used. If not otherwise mentioned, the aqueous phase was held in the syringe while the apolar phase was kept in a HEL-101-OS-20T cell from Hellma. The principle of the drop profile analysis is based on determination of the coordinates of a liquid drop from a video image and comparison of these coordinateswiththeoreticalprofilescalculatedfromtheGauss-Laplace equation.12 All the measurements were performed using the area mode control of an aqueous drop of 2.97 mm3. Interfacial Shear Rheology. The interfacial shear viscosity was measured with the ISR1 (SINTERFACE Technologies, Berlin) shear rheometer. The principle of these measurements is based on a biconical disk (measurement body) mounted at a torsion wire.13 The torsion wire is fixed to a deflection drive (a stepper motor). By an instantaneous movement of the torsion head, a torque is applied to the measurement body placed in the planar fluid interface of water-oil in a measuring vessel. A shear field is built up between the inner wall of the measuring vessel and the outer part of the measurement body (sharp edge of biconical disk). The angular displacement of the biconical disk is recorded by the instrument. After deflection, the pendulum performs damped harmonic oscillations with a damping factor R and a radian frequency β. The angular position of the measurement body is registered via a minilaser and a positionsensitive photosensor with an accuracy of 0.01°. The registered oscillation curve is then fitted by a model in order to determine the damping factor R and the radian frequency β of the torsion oscillation. From the difference between the measured values and those obtained for oscillation at a pure water-decane interface, the rheological parameters are calculated using the following formulas:
ηS ) 2HSIr(R - R0)
(1)
GS ) HSIr(R2 - R02 + β2 - β02)
(2)
where ηS is the interfacial shear coefficient of viscosity, GS is the interfacial shear modulus of rigidity, HS is the apparatus constant, depending on the shear field geometry, Ir is the moment of inertia of the measurement system, R is the damping factor, and β is the frequency of the torsion oscillation. Each 10 min, a measurement of interfacial shear rheology was performed with a duration of about 25 s per single oscillation measurement. The measurements were performed in a temperature controlled room at 20 °C using a deflection angle of 1.5°. A calibration measurement at the pure water-decane interface was used to estimate the values for R0 and β0. These measurements were carried out for more than 2 h in order to monitor the eventual adsorption or desorption of interfacially active impurities to or from the liquid-liquid interface. The nonaqueous phase must be fixed at the sharp edge of the biconical disk in order to prevent any wetting of the upper part of disk. After positioning the biconical disk, the aqueous phase and the disk are covered with the second fluid to a constant level. Exactly the same filling levels must be used for all measurements with interfacially active compounds as those for the reference measurement. The (12) Rotenberg, Y.; Boruvka, L.; Neumann, A. W. J. Colloid Interface Sci. 1983, 93, 169. (13) Kra¨gel, J.; Siegel, S.; Miller, R.; Born, M.; Schano, K. Colloids Surf., A 1994, 91, 169.
measurements were performed as a function of adsorption time with t ) 0 set when the filling procedure was finished. Ellipsometry. Ellipsometric measurements were performed with a Multiskop from Optrel, Berlin, Germany. The principle of this technique, the instrument used, and the procedure to calculate layer thickness and adsorbed amount have been described in detail elsewhere.14,15 For a thin, not light absorbing, plane-parallel, homogeneous, and isotropic layer of thickness δ , laser wavelength λ located at the interface between two liquid phases, the following equation holds:16
[
tan Ψei∆ ) tan Ψ0ei∆0 1 +
(
i
4πδ cos φ0 sin2 φ0n22M
λ(n22 - n02)(n02 sin2 φ0 - n22cos2 φ0)
)]
(3)
where ∆ and Ψ are the ellipsometric angles for the layer covering the interface. The parameters ∆0 and Ψ0 correspond to the interface without this layer, n1 is its refractive index, n0 and n2 are the refractive indices of ambient and substrate phases, respectively, φ0 is the angle of incidence, and M ) n02 + n22 - n12 - n02n22/n12. Using a numerical iteration procedure, the refractive index and the thickness of the adsorbed layer were calculated from the measured ellipsometric angles ∆, ∆0, Ψ, and Ψ0 via eq 3. The ellipsometric experiments were carried out in the following way. A clean Teflon vessel was carefully filled with freshly prepared lipase solution. The oil phase composed of diglyceride, monoglyceride, or fatty acid solution in decane was then gently poured above aqueous phase. After that, two light guides attached to the laser and detector arms of the ellipsometer were immersed into the upper oil phase so that incident light does not change its polarization state until reflection at a liquid-liquid interface under investigation. These light guides are cylindrical tubes made of stainless steel with the optical glass window on the bottom oriented perpendicularly to the propagation direction of the light, and they were introduced for the first time by Benjamins et al.17 The height of the ellipsometric table was then adjusted in such a manner that the level of the interface under investigation was set to reflect the beam slightly above the diaphragm on the detector arm. About 20 s later, the interface between the two liquids was cleaned by aspiration using a Pasteur pipet. Because of the small removal of both liquids due to aspiration, the interface level falls a bit and is now aligned to reflect the beam exactly through the middle pinhole on the diaphragm. The experiment was started a couple of seconds after the mechanical equilibration of the interface was attained. This procedure allowed thorough and fast cleaning of the investigated interface. Moreover, the loss of substance and any corresponding change in bulk concentration caused by cleaning were negligible, even at the lowest concentrations. All measurements were carried out at 23.0 ( 0.5 °C.
Results Interfacial Activity of Lipase from Rhizomucor miehei and of Products from Tricaprylin Hydrolysis. Figure 1a shows the interfacial activity of lipase from Rhizomucor miehei and of the amphiphilic products generated from hydrolysis of tricaprylin, that is, caprylic acid, monocaprylin, and dicaprylin, at a 0.1 M phosphate buffer pH 7.0-decane interface. The figure clearly shows that monocaprylin is the most interfacially active molecule (14) Miller, R.; Fainerman, V. B.; Makievski, A. V.; Kra¨gel, J.; Grigoriev, D. O.; Ravera, F.; Liggieri, L.; Kwok, D. Y.; Neumann, A. W. In Characterisation of water/oil interfaces, Encyclopaedic Handbook of Emulsion Technology; Sjo¨blom, J., Ed.; Marcel Dekker: New York, 2001; p 1. (15) Ducharme, D.; Tessier, A.; Leblanc, R. M. ReV. Sci. Instrum. 1987, 58, 571. (16) De Feijter, J. A.; Benjamins, J.; Veer, F. A. Biopolymers 1978, 17, 1759. (17) Benjamins, J. W.; Jo¨nsson, B.; Thuresson, K.; Nylander, T. Langmuir 2002, 18, 6437.
7402 Langmuir, Vol. 24, No. 14, 2008
Figure 1. Interfacial tension at a 0.1 M phosphate buffer pH 7.0-decane interface of (a) lipase from Rhizomucor miehei and the amphiphilic lipolysis products caprylic acid (C8), monocaprylin (MC8), and dicaprylin (DC8) or (b) binary mixtures of the lipolysis products at a fixed total concentration of 5.0 mM and with the interfacial tension values of only tricaprylin (TC8) and only lipase indicated at a 0.1 M phosphate buffer pH 7.0-decane interface. φ means molar ratio of the amphiphiles with MC8 to the left in the MC8+C8 and MC8+DC8 combinations and with DC8 to the left in the DC8+C8 combination.
generated in the lipolysis. It can also be seen that the monoglyceride gives considerably lower values of interfacial tension than the lipase. From an interfacial energy viewpoint, it is therefore reasonable to assume that a monoglyceride can expel a lipase from the interface provided that the monoglyceride is not a substrate for the enzyme.18 Under in ViVo conditions, a monoglyceride will coexist with other amphiphilic lipolysis products, in particular, diglyceride and fatty acid (or fatty acid salt). Figure 1b shows that the strong interfacial activity of the monoglyceride is retained also in combination with these two hydrolysis products. The interfacial tension values obtained with the combinations are very close to the values obtained with monoglyceride only, calculated on the amount of monoglyceride in the mixture. Effect of Addition of Sn-1/3 or Sn-2 Monoglyceride on the Free Energy of a Phosphate Buffer-Decane Interface Covered by Lipase Sn-1/3 and Sn-2 monoglycerides give the same interfacial tension (data not shown). However, when added to an interface already covered by a lipase having Sn-1,3 (18) Canioni, P.; Julien, R.; Tathelot, J.; Sarda, L. Biochimie 1976, 58, 751.
Reis et al.
Figure 2. Interfacial tension of the buffer/decane system with 3.3 × 10-5 M lipase from Rhizomucor miehei added followed by injection into the oil phase of (a) 5.0 × 10-3 M Sn-1/3 monocaprylin (Sn-1/3 MC8) or Sn-2 monocaprylin (Sn-2 MC8), or (b) 6.7 ×10-4 M Sn-1/3 monopalmitin (Sn-1/3 MC16) or Sn-2 monopalmitin (Sn-2 MC16).
regiospecificity, there is a major difference in time dependency with respect to the interfacial tension. This is illustrated in Figure 2 for monocaprylin (part a) and monopalmitin (part b). The monoglyceride is injected into the oil phase 1000 s after addition of the lipase. As can be seen, the Sn-2 monoglyceride reduces the interfacial tension to very low values, which do not change with time. The situation is different with the Sn-1/3 monoglyceride. Immediately after addition, the interfacial tension drops below the value of that of pure lipase but then rapidly recovers. The only reasonable explanation of this behavior is that the Sn1/3 monoglyceride, but not the Sn-2 monoglyceride, is rapidly hydrolyzed to the less interfacially active degradation products fatty acid and glycerol, which cannot compete with the lipase for a place at the interface. By comparing parts a and b of Figure 2 one can see that the dip in interfacial tension occurs during a shorter time scale for Sn-1/3 monopalmitin than for Sn-1/3 monocaprylin. Evidently, the lipase-catalyzed hydrolysis of monopalmitin occurs faster than that of monocaprylin. This is noteworthy in light of the fact that monocaprylin is the most interfacially active of the two and should therefore have a stronger driving force for the interface than monopalmitin. The latter is a relatively hydrophobic amphiphile that to a considerable extent partitions into the oil phase. The difference in the rate of increase of the interfacial tension found for the monoglycerides could be attributed either to a difference in specificity for the lipase or to
Lipase and Monoglyceride Competition at Interfaces
Figure 3. Interfacial tension of the buffer/decane system with 3.3 × 10-6 M lipase from Rhizomucor miehei added followed by injection into the oil phase of 6.7 × 10-4 M Sn-1/3 monopalmitin (Sn-1/3 MC16) or Sn-2 monopalmitin (Sn-2 MC16).
Figure 4. Interfacial tension of the buffer/decane system with 3.3 × 10-5 M lipase from Rhizomucor miehei added followed by injection into the oil phase of 1.3 ×-3 M Sn-1,3 dilaurin (Sn-1,3 DC12) or Sn-1,2 dilaurin (Sn-1,2 DC12).
a difference in partitioning of the in situ generated products. Similar results were obtained for lipoprotein lipase and pancreatic lipase in the presence of colipase (data not shown). The results shown in Figure 2 relate to events at an interface initially fully covered by the lipase. This may not always be realistic under in ViVo conditions. Figure 3 shows the effect on the interfacial tension when Sn-1/3 or Sn-2 monopalmitin is added to an interface that is only partly covered by the lipase. As expected, Sn-2 monopalmitin very rapidly takes over at the interface. The process is slow with Sn-1/3 monopalmitin, but the interfacial tension eventually attains the value characteristic of lipase, clearly as a result of lipase-catalyzed hydrolysis of the monoglyceride. Figure 4 gives further evidence of generated Sn-2 monoglyceride dominating the oil-water interface. Either Sn-1,2 or Sn1,3 dilaurin is added to the water phase of a phosphate-buffer/ decane system in which the interface is covered by the Sn-1,3 regiospecific lipase. Addition of the Sn-1,2 but not the Sn-1,3 diglyceride resulted in a lowering of the interfacial tension down to the value of pure monoglyceride.
Langmuir, Vol. 24, No. 14, 2008 7403
Figure 5. Interfacial tension of the buffer/decane system with 6.7 × 10-4 M Sn-1/3 monopalmitin (Sn-1/3 MC16) or Sn-2 monopalmitin (Sn-2 MC16) added followed by injection into the water phase of 3.3 × 10-5 M lipase from Rhizomucor miehei in the aqueous phase.
Effect of Addition of Lipase on the Free Energy of a Phosphate Buffer/Decane Interface Covered by Sn-1/3 or Sn-2 Monopalmitin Figure 5 shows the effect on the interfacial tension of adding a Sn-1,3 regiospecific lipase after 1000 s to the phosphate buffer/decane system to which either Sn-1/3 or Sn-2 monopalmitin has been preadded. As is seen from the figure, the interfacial tension attains approximately the same low value with both regioisomers. When the Sn-2 monoglyceride is covering the interface, the interfacial tension value is unaffected by the lipase addition because the enzyme is unable to catalyze cleavage of this monoglyceride due to its Sn-1,3 regiospecificity. With the Sn-1/3 monoglyceride at the interface, on the other hand, there is a steady increase in interfacial tension. This can only be explained by lipase-catalyzed hydrolysis of the monoglyceride into palmitic acid and glycerol, which are less interfacially active than the enzyme. Therefore, the interfacial tension increases to the value of pure lipase at the interface. Effect of Tricaprylin Hydrolysis Catalyzed by Lipase from Rhizomucor miehei or from Candida rugosa on the Free Energy of a Phosphate Buffer-Decane Interface Figure 6 shows the effect on the interfacial tension of the lipase-catalyzed hydrolysis of tricaprylin in a phosphate buffer/decane system. The lipase is either from Rhizomucor miehei, which is Sn-1,3 regiospecific and will yield Sn-2 monocaprylin together with caprylic acid, or from Candida rugosa, which is nonregiospecific and will give complete hydrolysis of tricaprylin into caprylic acid and glycerol. As seen in the figure, the reaction catalyzed by the lipase from Rhizomucor miehei results in a value of interfacial tension that is typical of that of a Sn-2 monocaprylincovered interface. The reaction catalyzed by the nonregiospecific Candida rugosa lipase gives an interfacial tension representative of that of a lipase-covered interface. The stable plateau value obtained from the hydrolysis catalyzed by the lipase from Rhizomucor miehei is an indication that the acyl group migration, that is, transition from Sn-2 monoglyceride to Sn-1/3 monoglyceride, is slow. Interfacial Dilational Rheology. The effect of adding Sn-2 monoglyceride on the elastic modulus of the phosphate buffer-decane interface covered by lipase from Rhizomucor miehei was studied. Figure 7 shows the elastic modulus as a function of oscillation frequency for the phosphate buffer-decane interface. As reference, the same experiment was performed with a noncatalytically active protein, β-lactoglobulin (BLG). The
7404 Langmuir, Vol. 24, No. 14, 2008
Reis et al.
Figure 6. Change in interfacial tension of the buffer/decane system containing 6.1 × 10-3 M tricaprylin on addition of 3.3 × 10-5 M lipase from either Rhizomucor miehei (Lipase R.m.) or Candida rugosa (Lipase C.r.).
Figure 7. Elastic modulus versus oscillation frequency for the buffer-decane interface covered with 3.3 × 10-5 M Rhizomucor miehei lipase, 7.14 × 10-4 M β-lactoglobulin (BLG), 6.4 × 10-4 M Sn-2 monopalmitin (Sn-2 MC16), lipase + Sn-2 monopalmitin, or BLG + Sn-2 monopalmitin.
interface is covered by either the lipase or BLG, and the concentrations of the proteins are chosen such that the two systems have the same interfacial tension, 15 mN/m. As can be seen from the figure, the elastic moduli of the two interfaces are very different. Whereas the BLG-covered interface has a high elastic modulus that increases with the oscillation frequency, the lipasecovered interface has values of the same order as the interface covered by only Sn-2 monopalmitin. Addition of Sn-2 monopalmitin to the protein-containing systems results in a drastic reduction in the elastic modulus for the BLG system and a small reduction for the lipase system, in both cases most likely an effect of the protein being expelled from the interface by the more surface-active Sn-2 monoglyceride. Interfacial Shear Rheology. With tensiometry, one follows the change in surface free energy, but the technique does not per se give information about what molecules are present at the
interface. Interfacial shear rheology gives complementary information because it can distinguish between low and high molecular weight amphiphilic compounds in the interfacial layer. The torsion pendulum technique, which has been used in this work, gives information about the stiffness of the interfacial layer. By measuring the variation of oscillation amplitude with time, a change in the rheological characteristics of this layer can be detected. Figure 8 shows the response to shear deformation of the phosphate buffer-decane interface covered by either a crude preparation of lipase from porcine pancreas or BLG at concentrations corresponding to full surface coverage. Measurements were made immediately after the formation of the interface, after 30 min rest, and after 60 min rest. With the crude lipase preparation at the interface, there is some initial oscillation but it is rapidly damped. The experiment after 60 min rest shows a lower oscillation amplitude as a result of an increased stiffness of the interfacial layer. In a recent publication,19 we showed that lipase from Rhizomucor miehei behaves like a small molecular weight surfactant at the interface (even when the interface is saturated with the enzyme). Therefore, the relatively high stiffness seen in the interfacial shear rheology experiments for the crude preparation of pancreatic lipase is most likely due to either anchoring of the lipase to the interface by the colipase or the presence of other proteins in the lipase preparation. The purpose of using pancreatic lipase instead of a pure lipase that does not require a colipase is to assess the interfacial behavior of physiologically relevant enzyme preparations. With BLG at the interface, there is no oscillation whatsoever. The pendulum is completely damped. Only a creeping of the disk to a new angular position is observed. Figure 9 shows curves of the change in the torsion angle with time for Sn-2 monopalmitin added to the phosphate buffer/decane system. The damping pattern is the same regardless of the time between addition of the monoglyceride and the experiment. Very (19) Reis, P.; Miller, R.; Kra¨gel, J.; Leser, M. E.; Fainerman, V. B.; Watzke, H.; Holmberg, K. Langmuir 2008, 24, 6812.
Lipase and Monoglyceride Competition at Interfaces
Langmuir, Vol. 24, No. 14, 2008 7405
Figure 10. Variation of the torsion angle with time after formation of the buffer-decane interface with the water phase containing 10 mg/mL lipase from porcine pancreas and 6.6 × 10-4 M Sn-1/3 monopalmitin.
Figure 8. Variation of the torsion angle with time after formation of the buffer-decane interface with the water phase containing (a) a 10 mg/ mL solution of lipase from porcine pancreas and (b) 7.14 × 10-4 M β-lactoglobulin. Figure 11. Variation of the torsion angle with time after formation of the buffer-decane interface with the water phase containing 10 mg/mL lipase from porcine pancreas and 6.6 × 10-4 M palmitic acid.
Figure 9. Variation of the torsion angle with time after formation of the buffer-decane interface with the water phase containing 6.6 × 10-4 M Sn-2 monopalmitin.
similar curves were obtained when Sn-2 monopalmitin was replaced by either Sn-1/3 monopalmitin or palmitic acid, and the curves resemble those obtained in a pure phosphate buffer/decane system. Thus, even if the monoglyceride is a very interfacially
active amphiphile, which implies tight packing, the experiment indicates that there is no increased interfacial stiffness. The effect of added Sn-1/3 monopalmitin on the interfacial shear rheology behavior of pancreatic lipase was studied, and the results are shown in Figure 10. Monopalmitin will rapidly be hydrolyzed to palmitic acid and glycerol. By comparing the curves of Figure 10 with those of Figure 8a, one can see that the system with lipase and hydrolysis products differs from that with only lipase. The curve from the initial run (0 min) is identical to that from the 0 min run of Figure 8a. The 30 and 60 min runs are very different, however. These curves show no oscillations. Most likely, the strong damping is caused by the generated fatty acid, as can be seen from Figure 11, which shows the behavior of a combination of lipase from porcine pancreas and palmitic acid. The variations of the torsion angle with time for palmitic acid at the buffer-decane interface at pH 7.0 are similar to those of a bare interface (data not shown). Therefore, the hypothesis of liquid crystalline phase formation was not considered. Figure 12 shows the corresponding curves for the addition of Sn-2 monopalmitin to the same two-phase system with the Sn1,3 regiospecific lipase from porcine pancreas covering the interface. The damping pattern is identical to that of Figure 9.
7406 Langmuir, Vol. 24, No. 14, 2008
Figure 12. Variation of the torsion angle with time after formation of the buffer-decane interface with the water phase containing 10 mg/mL lipase from porcine pancreas and 6.6 × 10-4 M Sn-2 monopalmitin.
Reis et al.
Lipase has been found to undergo slow secondary structure changes during the first stages of adsorption.20 The isoelectric point of lipase is about 3.5, that is, it is negatively charged at pH 7, which is the pH used in the experiments. It is known that OH- ions accumulate at an oil-water interface, rendering it negatively charged.21 Therefore, there will be an electrostatic repulsion between the lipase and the oil-water interface working against extensive adsorption.22 As a result, the adsorbed layer becomes loose with a protein concentration not very much higher than the concentration in the bulk phase. This leads to an apparent enhancement of the layer thickness in Figure 13. The obtained ellipsometric thickness is not strictly the thickness of the adsorbed layer but rather the thickness of the uppermost region of aqueous phase, where the lipase concentration is higher than that in the bulk. However, water is the dominant component of this layer. To simplify the evaluation of the ellipsometric data, we used a fixed value for the refractive index of the adsorbed layer, n1 ) 1.375, which is quite close to the water value. As can be seen, the thickness decreases somewhat when Sn1/3 monopalmitin is added and the value obtained is exactly the same as that obtained by addition of the same molar concentration of palmitic acid. The reduction in thickness is probably due to changes in conformation leading to a denser layer, as was discussed in connection to the interfacial shear rheology experiments. When Sn-2 monopalmitin is added, the thickness increases. The monoglyceride will take over the very interface but the expelled lipase evidently accumulates in the interfacial region.
Discussion
Figure 13. Effect of monopalmitin (either Sn-1/3 MC16 or Sn-2 MC16) or of palmitic acid (C16) on the interfacial thickness of a lipase-covered interface between the buffer and decane. The lipase was from Rhizomucor miehei and had been introduced as a 3.3 × 10-5 M water solution.
The result is further proof of the Sn-2 monoglyceride rapidly replacing the lipase from the interface. The situation at the interface is stable during the time of the experiment, 60 min, as judged from the identical damping curves. Interfacial Layer Thickness. Ellipsometry was used to investigate whether the lipase that is expelled from the oil-water interface by the Sn-2 monoglyceride goes out into the bulk or remains in the interfacial region. The same phosphate buffer/ decane system was used as in the previous experiments, and the Sn-1,3 regiospecific lipase from Rhizomucor miehei was introduced in order to cover the interface with the enzyme. Either Sn-1/3 monopalmitin or Sn-2 monopalmitin was subsequently added. From the previous experiments, we know that whereas Sn-1/3 monopalmitin will be rapidly hydrolyzed to palmitic acid and glycerol, Sn-2 monoglyceride will not be degraded. We also know that the Sn-2 monopalmitin, but not the generated palmitic acid, will replace the lipase at the interface. The interfacial layer thickness of the different systems as determined by ellipsometry is shown in Figure 13.
The results from the tensiometry measurements and from the interfacial shear rheology measurements point in the same direction and show conclusively that out of the hydrolysis products formed in fat digestion only the monoglyceride has enough interfacial activity to replace the lipase from the interface. Lipases are either Sn-1,3 regiospecific or nonregiospecific. Triglyceride hydrolysis catalyzed by a Sn-1,3 regiospecific lipase will yield Sn-2 monoglyceride and fatty acid, while hydrolysis catalyzed by a nonregiospecific lipase will give fatty acid and glycerol. Thus, if lipolysis occurs with a Sn-1,3 regiospecific lipase, but not with a nonspecific lipase, a reaction product will form that is so interfacially active that the lipase is expelled from the interface. For this to occur, the rate of the acyl group migration, that is, transformation of the Sn-2 monoglyceride into the more thermodynamically favorable Sn-1/3 monoglyceride, must be slow. We have shown that, under the conditions used here, this seems to be the case. Since lipases are water-soluble enzymes and their main substrate, the triglyceride, is not, they always operate at an oil-water interface. If they are expelled from the interface, lipolysis is likely to cease. For this to occur during fat digestion, that is, through replacement by generated 2-monoglyceride, the lipase needs to be Sn-1,3 regiospecific. The enzymes that operate in the gastrointestinal system, such as pancreatic lipase, exhibit Sn-1,3 regiospecificity. In this work, we have used pancreatic lipase but also another Sn-1,3 regiospecific lipase, from Rhizomucor miehei, to demonstrate that the Sn-2 monoglyceride formed takes over at the interface, expelling (20) Noinville, S.; Revault, M.; Baron, M.; Tiss, A.; Yapoudjian, S.; Ivanova, M.; Verger, R. Biophys. J. 2002, 82, 2709. (21) Marinova, K. G.; Alargova, R. G.; Denkov, N. D.; Velev, O. D.; Petsev, D. N.; Ivanov, I. B.; Borwankar, R. P. Langmuir 1996, 12, 2045. (22) Campos, J.; Eskilsson, K.; Nylander, T.; Svendsen, A. Colloids Surf., B 2002, 26, 172.
Lipase and Monoglyceride Competition at Interfaces
the lipase from what can be regarded as the reaction zone. We have also seen that β-lactoglobulin, which lacks enzymatic activity, was desorbed equally well by both monoglyceride isomers (either Sn-1/3 or Sn-2). In a separate communication,11 where we have used the TNO gastrointestinal model (TIM), a model for the gastrointestinal system, we have shown that fat digestion is discontinued when Sn-2 monoglyceride is added to the model meal. Thus, this can be regarded as a self-regulating process that prevents excessive digestion of the available triglycerides. The results also suggest a way to minimize the energy uptake of fat food by adding Sn-2 monoglyceride as a food additive. In order to support our findings that Sn-2 monoglycerides prevent lipolysis, we used a nonregiospecific lipase from Candida rugosa as a control. Tensiometry experiments showed that this lipase remained at the interface after fat digestion. The fatty acid (or fatty acid salt) formed in the complete hydrolysis of the triglyceride did not have enough interfacial activity to expel the lipase. Previous articles have suggested that nonfood grade nonionic surfactants such as Brij 35 and Triton X-100 can displace lipase and colipase from the interface.16 This is likely in view of our results, but in this work we have not investigated the influence of external surfactants but concentrated on the self-regulatory mechanism, where lipase generates surface-active molecules, which compete with the enzyme for the interface. A deeper insight into the interfacial competition between proteins and monoglycerides was provided by the rheological experiments. When the elastic modulus is assessed for a lipase and for BLG as a function of the frequency of oscillation, one can observe that the elastic modulus is much lower for the lipase than for BLG. Actually, the value recorded for the lipase is closer to the value obtained with Sn-2 monopalmitin than to that obtained with BLG. This difference in interfacial rheology between the lipase and BLG indicates that the lipase remains in an unperturbed conformation at the interface, which is likely to be a requirement for retention of the biological activity. The interfacial shear rheology measurements confirm that a crude mixture of pancreatic lipase and colipase is displaced from the interface by Sn-2 monoglycerides. Therefore, the capacity of the colipase to anchor the lipase at the interface is limited. The same result was obtained with BLG (data not shown). Our results support the model by Haiker et al.,23 suggesting that pancreatic lipase does not remain irreversibly adsorbed at the oil-water interface. It may exchange between the surfaces of oil droplets in an emulsion. As discussed previously, Sn-1/3 monoglycerides do not displace lipase from the interface. Instead, the in situ generation of free fatty acids increased the stiffness of the protein layer at the interface. This result may be attributed to the formation of lipase-fatty acid complexes at the oil-water boundary.24 Previous studies have shown that monoglycerides (monopalmitin and monoolein) and proteins (β-casein and caseinate) may form heterogeneously mixed films at the air-water interface.25,26 However, the type of interaction that is developed between a monoglyceride and a protein will vary for different monoglyceride-protein combinations. As an example, it has been observed that interactions between proteins and monoglyc(23) Haiker, H.; Lengsfeld, H.; Hadva´ry, P.; Carrie`re, F. Biochim. Biophys. Acta 2004, 1682, 72. (24) Bengtsson, G.; Olivercrona, T. Eur. J. Biochem. 1980, 106, 557. (25) Rodriguez Patino, J. M.; Carrera, C. S.; Rodriguez Nin˜o, M. J. Agric. Food Chem. 1999, 47, 4998. (26) Rodriguez Patino, J. M.; Carrera, C. S.; Rodriguez Nin˜o, M. Langmuir 1999, 15, 4777.
Langmuir, Vol. 24, No. 14, 2008 7407
erides are stronger for globular proteins, such as BLG, than for disordered proteins, such as β-casein. In fact, BLG is known to be a lipid binding protein.27 Lipase, being a globular protein, should then interact strongly with monoglycerides. Our ellipsometry results show that pancreatic lipase remains in a sublayer below the oil-water interface, which is saturated with Sn-2 monoglycerides. The concept of proteins residing underneath a monoglyceride film, with which they interact by van der Waals forces and/or by acid-base interactions involving the headgroup layer of the amphiphiles, has been suggested earlier.28–30 We have recently shown that a lipase can also adsorb at hydrophilic surfaces31 but to a lesser extent than at hydrophobic ones. From our results, it is reasonable to assume that during the digestion of triglycerides, lipase will be displaced from the interface by Sn-2 monoglycerides accumulating at the interface. However, the enzymatic activity is not entirely hampered by the exclusion from the interface. In a separate communication, we show that provided that the interface is populated by Sn-2 monoglycerides and free fatty acids, the enzyme continues to catalyze esterification reactions.32 In in ViVo systems, the in situ generated Sn-2 monoglycerides are solubilized by bile salts from the oil-water interface and transported across the brush border of the gastrointestinal system. This delivery mechanism provides a re-establishment of the solubilization capacity of bile salts and a high yield of fat absorption. This process does not occur in the in Vitro TIM model, and therefore, the bile salts remain saturated with Sn-2 monoglycerides. Therefore, the accumulation of this reaction product at the interface controls the lipolysis. It has been reported that, at least during the initial stages of fat digestion, the rate of lipolysis significantly exceeds the capacity to incorporate the resulting fatty acids and monoglycerides into mixed bile salt micelles.33 These polar lipids accumulate at their site of origin, the oil-water interface, leading to a self-regulation mechanism. Consequently, the high efficiency of fat digestion in in ViVo systems is attributed to the transport mechanisms of interfacially active molecules from the oil-water interface and to further delivery of these molecules at the brush border cells. Any mechanism that affects the bile salt solubilization capacity (e.g., ionic strength) should then affect the rate and extent of fat digestion.
Concluding Remarks This paper demonstrates, by combining results from tensiometry, interfacial rheology, and ellipsometry, that Sn-2 monoglycerides expel Sn-1,3 regiospecific lipases from the oil-water interface. The lipase appears to form a sublayer in the water phase, located just beneath the monoglyceride-covered interface. Since the triglycerides are present in, or constitute, the apolar phase of the system and since the lipases involved in lipolysis all exhibit Sn-1,3 regiospecificity, the removal of the enzyme from the reaction zone can be seen as a self-regulatory mechanism that controls the extent of fat digestion. LA800531Y (27) Papiz, M. Z.; Sawyer, L.; Eliopoulos, E. E.; North, A. C.; Findlay, J. B.; Sivaprasadarao, R.; Jones, T. A.; Newcomer, M. E.; Kraulis, P. J. Nature 1986, 324, 383. (28) Li, J. B.; Kra¨gel, J.; Makievski, A. V.; Fainermann, V. B.; Miller, R.; Mo¨hwald, H. Colloids Surf., A 1998, 142, 355. (29) Cornec, M.; Narismhan, G. Langmuir 2000, 16, 1216. (30) Rodriguez Patino, J. M.; Rodriguez Nin˜o, M.; Sanchez, C. C.; Fernandez, M. C. Langmuir 2001, 17, 7545. (31) Reis, P.; Holmberg, K.; Debeche, T.; Folmer, B.; Fauconnot, L.; Watzke, H. Langmuir 2006, 22, 8169. (32) Reis, P.; Holmberg, K.; Miller, R.; Leser, M.; Watzke, H. Lipase catalyzed reactions at interfaces of two-phase systems and microemulsions. To be submitted. (33) Carey, M.; Hernell, O. Semin. Gastrointest. Dis. 1992, 3, 189.
