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Adsorption of Bile Salts and Pancreatic Colipase and Lipase onto Digalactosyldiacylglycerol and Dipalmitoylphosphatidylcholine Monolayers Boon-Seang Chu, A. Patrick Gunning, Gillian T. Rich, Mike J. Ridout, Richard M. Faulks, Martin S. J. Wickham, Victor J. Morris, and Peter J. Wilde* Institute of Food Research, Norwich Research Park, Colney Lane, Norwich NR4 7UA, U.K. Received January 5, 2010. Revised Manuscript Received February 23, 2010 It is increasingly recognized that changes in the composition of the oil-water interface can markedly affect pancreatic lipase adsorption and function. To understand interfacial mechanisms determining lipase activity, we investigated the adsorption behavior of bile salts and pancreatic colipase and lipase onto digalactosyldiacylglycerol (DGDG) and dipalmitoylphosphatidylcholine (DPPC) monolayers at the air-water interface. The results from Langmuir trough and pendant drop experiments showed that a DGDG interface was more resistant to the adsorption of bile salts, colipase, and lipase compared to that of DPPC. Atomic force microscopy (AFM) images showed that the adsorption of bile salts into a DPPC monolayer decreased the size of the liquid condensed (LC) domains while there was no visible topographical change for DGDG systems. The results also showed that colipase and lipase adsorbed exclusively onto the mixed DPPC-bile salt regions and not the DPPC condensed phase. When the colipase and lipase were in excess, they fully covered the mixed DPPC-bile salt regions. However, the colipase and lipase coverage on the mixed DGDG-bile salt monolayer was incomplete and discontinuous. It was postulated that bile salts adsorbed into the DPPC monolayers filling the gaps between the lipid headgroups and spacing out the lipid molecules, making the lipid hydrocarbon tails more exposed to the surface. This created hydrophobic patches suitable for the binding of colipase and lipase. In contrast, bile salts adsorbed less easily into the DGDG monolayer because DGDG has a larger headgroup, which has strong intermolecular interactions and the ability to adopt different orientations at the interface. Thus, there are fewer hydrophobic patches that are of sufficient size to accommodate the colipase on the mixed DGDG-bile salt monolayer compared to the mixed DPPC-bile salt regions. The results from this work have reinforced the hypothesis that the interfacial molecular packing of lipids at the oil-water interface influences the adsorption of bile salts, colipase, and lipase, which in turn impacts the rate of lipolysis.
Introduction The digestion of dietary fat requires the hydrolysis of triacylglycerols to monoacylglycerols and fatty acids by gastric and pancreatic lipases, with 70-90% of the lipolysis taking place in the small intestine.1 Lipolysis occurs in the presence of various surface-active substances with a range of charge, structure, and surface activity originating from food, its digestion products, and gastric/pancreatic/hepatic secretions.2 Since lipolysis requires close proximity of lipase to the lipid substrate at the oil-water interface, the presence of these surface-active substances at the interface is thought to influence the interfacial reaction. For example, ovalbumin and β-lactoglobulin adsorbed onto oil-inwater emulsion surfaces have been shown to inhibit pancreatic lipase activity.3,4 The inhibitory effects are attributed to the ability of the amphiphiles to prevent direct contact between the lipase and lipid substrates. In the duodenum, secreted bile salts contain highly surface-active components (bile acids) which can displace other surface-active molecules from the emulsified lipid surfaces, but the accumulation of the negatively charged bile salts at the *To whom correspondence should be addressed: Tel þ44 (0)1603 255258; Fax þ44 (0)1603 507723; e-mail
[email protected].
(1) Fave, G.; Coste, T. C.; Armand, M. Cell. Mol. Biol. 2004, 50, 815. (2) Lowe, M. E. J. Lipid Res. 2002, 43, 2007. (3) Ivanova, M.; Panaiotov, I.; Bois, A. G.; Gargouri, Y.; Verger, R. J. Colloid Interface Sci. 1990, 136, 363. (4) Mandalari, G.; Adel-Patient, K.; Barkholt, V.; Baro, C.; Bennett, L.; Bublin, M.; Gaier, S.; Graser, G.; Ladics, G. S.; Mierzejewska, D.; Vassilopoulou, E.; Vissers, Y. M.; Zuidmeer, L.; Rigby, N. M.; Salt, L.; Defernez, M.; Mulholland, F.; Mackie, A. R.; Wickham, M. S. J.; Mills, E. N. C. Regul. Toxicol. Pharmacol. 2009, 55, 372. (5) Borgstr€om, B. J. Lipid Res. 1975, 16, 411. (6) Momsen, W. E.; Brockman, H. L. J. Biol. Chem. 1976, 251, 378.
9782 DOI: 10.1021/la1000446
interface also hinders the adsorption of the pancreatic lipase.5,6 This lipase inhibitory effect of bile salts is reversed by the presence of colipase which binds avidly to a bile salt-dominated interface and anchors pancreatic lipase to the interface via the formation of a specific 1:1 colipase-lipase complex.7 Thus, the adsorption of bile salts and colipase act together to regulate pancreatic lipase activity through their combined effects on the interfacial activation of lipase on the surfaces of emulsified lipid substrates. The adsorption of bile salts onto the lipid emulsion surfaces has been monitored in vitro by examining changes in the surface charge of oil emulsion droplets as a function of bile salt concentration.8,9 Bile salts have also been shown recently to displace protein networks in a very similar fashion and at similar concentrations from both the air-water and oil-water interfaces10 through an orogenic mechanism originally demonstrated using conventional surfactants.11-15 The incorporation of bile salts into (7) Donner, J.; Spink, C. H.; B€orgstrom, B.; Sjoholm, I. Biochemistry 1976, 15, 5413. (8) Wickham, M.; Garrood, M.; Leney, J.; Wilson, P. D. G.; Fillery-Travis, A. J. Lipid Res. 1998, 39, 623–632. (9) Chu, B. S.; Rich, G. T.; Ridout, M. J.; Faulks, R. M.; Wickham, M. S. J.; Wilde, P. J. Langmuir 2009, 25, 9352. (10) Maldonado-Valderrama, J.; Woodward, N. C.; Gunning, A. P.; Ridout, M. J.; Husband, F. A.; Mackie, A. R.; Morris, V. J.; Wilde, P. J. Langmuir 2008, 24, 6759. (11) Mackie, A. R.; Gunning, A. P.; Wilde, P. J.; Morris, V. J. J. Colloid Interface Sci. 1999, 210, 157. (12) Mackie, A. R.; Gunning, A. P.; Wilde, P. J.; Morris, V. J. Langmuir 2000, 16, 2242. (13) Mackie, A. R.; Gunning, A. P.; Wilde, P. J.; Morris, V. J. Langmuir 2000, 16, 8176. (14) Morris, V. J.; Gunning, A. P. Soft Matter 2008, 4, 943. (15) Woodward, N. C.; Gunning, A. P.; Mackie, A. R.; Wilde, P. J.; Morris, V. J. Langmuir 2009, 25, 6739.
Published on Web 03/11/2010
Langmuir 2010, 26(12), 9782–9793
Chu et al.
the interface promoted the disruption and breakup of the packing and structure of the original amphiphiles adsorbed at the interface.10,12-15 This breakup allows the bile salts to form clusters, which not only is of importance for the colipase and lipase binding to the interface but also allows the lipase access to its substrates for lipolysis.8 Another key event during duodenal lipolysis is the adsorption of colipase and lipase to the emulsion droplet surface. Colipase is a small, amphiphilic wedge-shaped protein with a “three-finger” topology.16,17 Previous studies have shown that colipase requires about 145 A˚2 of hydrophobic area for adsorption to take place even though it occupies about 500 A˚2 when fully adsorbed onto the surface.18,19 Following adsorption, colipase is thought to act as a nucleating center for substrate molecules at the interface to enable lipase to adsorb in the catalytically efficient conformation.18 It is not known with certainty if colipase adsorbs to the emulsion surfaces alone or as a complex with pancreatic lipase.18,20 However, crystallographic studies have shown that colipase binds to the noncatalytic C-terminal domain of pancreatic lipase and exposes the hydrophobic tips of its fingers at the opposite side of its lipase-binding site in bulk solution.16,17,21 In the active conformation, these hydrophobic tips help to bring the catalytic N-terminal domain of pancreatic lipase into close contact with the interface where the lipase lid domain adopts the open conformation22 and colipase interacts with the lipase to form a lipid-water interface binding site which is more than 50 A˚ in length with an area of about 900 A˚2.16,17,21 Indeed, the adsorption of colipase, and therefore lipase, is dependent on interfacial lipid composition and lateral packing and demonstrates a clear specificity for interfaces containing lipase substrates.18,23,24 Monolayer studies have shown that colipase adsorption is triggered at a substrate mole fraction below its so-called putative percolation threshold so that lipolysis can also occur at a lower substrate mole fraction at the interface.18,25 Once lipolysis is initiated on the emulsion surfaces, more substrate becomes available to lipase and the lipolysis proceeds autocatalytically.18 The work presented here aims to understand the influence of DGDG and DPPC interfacial behaviors on the adsorption of bile salts, colipase, and lipase at the air-water interface. The aim is to demonstrate the potential for the use of DGDG for moderating lipid digestion which could be used as a strategy for controlling appetite.26 DGDG, an uncharged surface-active lipid found in thylakoid and amyloplast membrane systems of plants,27,28 consists of a large digalactosyl headgroup (Figure 1A) and two usually unsaturated fatty acyl chains. In contrast, DPPC has a smaller headgroup (Figure 1B) and saturated dipalmitoyl chains. (16) Egloff, M.-P.; Marguet, F.; Buono, G.; Verger, R.; Cambillau, C.; van Tilbeurgh, H. Biochemistry 1995, 34, 2751. (17) Egloff, M.-P.; Sarda, L.; Verger, R.; Cambillau, C.; van Tilbeurgh, H. Protein Sci. 1995, 4, 44. (18) Sugar, I. P.; Mizuno, N. K.; Momsen, M. M.; Brockman, H. L. Biophys. J. 2001, 81, 3387. (19) Sugar, I. P.; Mizuno, N. K.; Brockman, H. L. Biophys. J. 2005, 89, 3997. (20) Allouche, M.; Castano, S.; Colin, D.; Desbat, B.; Kerfelec, B. Biochemistry 2007, 46, 15188. (21) van Tilbeurgh, H.; Egloff, M.-P.; Martinez, C.; Rugani, N.; Verger, R.; Cambillau, C. Nature 1993, 362, 814. (22) Bezzine, S.; Ferrato, F.; Ivanova, M. G.; Lopez, V.; Verger, R.; Carriere, F. Biochemistry 1999, 38, 5499. (23) Mizuno, N. K.; Smaby, J. M.; Cunningham, B. A.; Momsen, M. M.; Brockman, H. L. Langmuir 2003, 19, 1802. (24) Momsen, M. M.; Dahim, M.; Brockman, H. L. Biochemistry 1997, 36, 10073. (25) Brockman, H. Biochimie 2000, 82, 987. (26) Maljaars, P. J. W.; Peters, H. P. F.; Mela, D. J.; Masclee, A. M. Physiol. Behav. 2008, 95, 271. (27) Sastry, P. Adv. Lipid Res. 1974, 12, 251. (28) D€ormann, P.; Benning, C. Trends Plant Sci. 2002, 7, 112.
Langmuir 2010, 26(12), 9782–9793
Article
Figure 1. Chemical structure of (A) digalactosyldiacylglycerol and (B) dipalmitoylphosphatidylcholine. R1 and R2 are fatty acyl chains, and R3 is palmitoyl chain.
Incorporation of DGDG into olive oil emulsions has been shown to slow down the rate of lipolysis.9 Here we report results from pendant drop and Langmuir trough experiments with adsorbed and spread lipid monolayers, respectively. The experiments were designed to study the adsorption kinetics of bile salts, colipase, and lipase into the air-water interface and determine how the lipids’ molecular orientations and intermolecular reactions influence the results. The DGDG and DPPC monolayers were characterized by plotting their surface pressure (π)-area (A) isotherms. The monolayers, following the adsorption of bile salts, colipase, and lipase, were transferred from the air-water interface onto mica using the Langmuir-Blodgett (LB) technique and then visualized using AFM.11 This work provides interesting new insights into the basic mechanisms likely to occur during the adsorption of bile salts and enzymes onto the DGDG and DPPC monolayers at the air-water interface. The results provide a rational basis for understanding how interfacial packing affects lipolysis and so should be applicable in general terms to the oil-water interface in oil-in-water emulsions. One of the main reasons we are using an air-water interface is because it is not possible to perform these AFM experiments at the oil-water interface. To image the interfaces, we have to use solvent to remove the oil phase from the Langmuir-Blodgett film. This has been possible for previous work on proteins due to their poor solubility in solvent. However, the solvents will remove the polar lipids we are studying from the LB film, thus destroying the structure. As stated above, we have compared the adsorption and action of bile salts at both the air-water and oil-water interfaces9,10 and feel that under these conditions carefully designed experiments at the air-water interface provide useful information regarding the observed differences between DGDG and DPPC in regulating the adsorption of amphiphiles into the interface.9
Materials and Methods Materials. The solution of DGDG (>98% purity, from spinach leaf) in chloroform-methanol was purchased from Lipid Products (Surrey, UK) and used without further purification. 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC, >99% purity), highly purified porcine pancreatic lipase (Type VI-S, activity g20 000 units/mg protein), essentially salt-free porcine pancreas colipase, sodium glycodeoxycholate (NaGDC, >97% purity), and sodium taurocholate (NaTC, >95% purity) were obtained from Sigma-Aldrich Ltd. (Gillingham, UK). Ultrapure water (resistivity 18.2 MΩ/cm, surface tension 72.6 ( 0.5 mN/m at 20 °C) was produced using an Elga Elgastat UHQ water purification system (Elga Process Water, Marlow, UK). All glassware was washed with 10% Micro-90 cleaning solution and exhaustively rinsed with tap water, isopropanol, tap water, DOI: 10.1021/la1000446
9783
Article deionized water, and ultrapure water in sequence. All other chemicals used were of analytical grades. π-A Isotherms. A polytetrafluoroethylene (PTFE) Langmuir trough of 255 112 16 mm with a volume of 450 mL and equipped with one fixed and one computer-controlled movable barrier was used for plotting the π-A isotherms of DGDG and DPPC. The clean PTFE trough was filled with ultrapure water, and the temperature was set at 37.0 ( 0.5 °C with the aid of an external heated water bath. The surface pressure was measured using a wetted ground glass Wilhelmy plate (0.19 mm thickness and 23.88 mm width). The absence of surface-active contaminants in the subphase was verified by the measurement of surface tension and confirmed with a π-A isotherm of the water that did not change by more than 0.5 mN/m over the entire compression range. Aliquots of DGDG or DPPC (0.1 mg/mL in 1:1 v/v chloroform-methanol solution) were carefully spread at the interface at a low surface density and negligible surface pressure. After evaporation of the solvent and equilibration of the monolayer (ca. 15 min), the lipid monolayer was compressed using the movable barrier operated at a rate of 0.85 mm/s, and the corresponding change in surface pressure was recorded. Preparation of LB Films. DGDG and DPPC LB films were prepared as described previously.29 A DGDG or DPPC monolayer was spread on water using the same protocol described in the previous section. Water was used as the subphase as the buffer salts would have interfered with the AFM imaging. Compression isotherms showed very little difference in the presence of buffer or water (results not shown). The monolayer was compressed to reach the required surface pressure. After the surface pressure stabilized (ca. 30 min), LB films were produced by lowering a freshly cleaved piece of mica sheet perpendicularly down through the interface and then pulling it out again.11 Surface pressure was monitored during the dip and showed that the monolayer was only transferred onto the mica on the upward stroke. Mica was used as a supporting solid substrate for LB film deposition because it is atomically flat over the hundred nanometer scale and can be easily cleaved to form a clean flat substrate. The mounted piece of mica was driven at a constant rate of about 0.15 mm/s so that there was virtually no excess water left on the mica as it dewetted when coming out from the subphase. Nevertheless, the LB films were dried in air at room temperature before they were observed under AFM. These experiments were not possible at the oil-water interface, as removal of the oil phase with solvents prior to AFM imaging would also remove the polar lipids from the LB film. Bile Salt Adsorption into Lipid Monolayers. A lipid monolayer was compressed to a defined initial surface pressure of 15.0 ( 1.0 mN/m. After the initial surface pressure became stable, 0.1 mL of 80 mM bile salt solution was carefully injected into the subphase from the outside of the barriers of the trough, resulting in a bile salt concentration of about 0.018 mM in the subphase. The bile salt solution contained a mixture of NaTC and NaGDC in a proportion of 52.7 and 47.3 mol %, respectively, in order to mimic the hydrophobicity of bile salts in human bile.30,31 Injection of the bile salt solution increased the surface pressure, indicating the adsorption of bile salts into the lipid monolayer. Eight successive additions of bile salt solution into the subphase were made, each comprising 0.1 mL of 80 mM bile salt solution, to progressively increase the surface pressure which was monitored by means of the Wilhelmy plate. The interfacial structure was transferred onto freshly cleaved mica substrate using the LB technique after the first, third, and eighth additions of bile salt solution. The bile salt adsorption experiments were also performed at a higher surface pressure, similar to that at an oil-water (29) Lucero, A.; Rodrı´ guez Ni~no, M. R.; Gunning, A. P.; Morris, V. J.; Wilde, P. J.; Rodrı´ guez Patino, J. M. J. Phys. Chem. B 2008, 112, 7651. (30) Armstrong, M. J.; Carey, M. C. J. Lipid Res. 1982, 23, 70. (31) Staggers, J. E.; Hernell, O.; Stafford, R. J.; Carey, M. C. Biochemistry 1990, 29, 2028.
9784 DOI: 10.1021/la1000446
Chu et al. emulsion interface in the presence of bile salts at a physiologically relevant concentration.9,10 For this purpose, the initial surface pressure of the lipid monolayer was set at 25.0 ( 1.0 mN/m and the bile salt concentration in the subphase was brought to 9.7 mM in one single injection. The change in surface pressure upon the bile salt addition was monitored and recorded. For comparative purposes, the surface pressure induced at a clean air-water interface by bile salts at concentrations of 0.018 and 9.7 mM was also measured. Lipase and Colipase Adsorption. DGDG and DPPC LB films with adsorbed bile salts, colipase, and lipase were also studied. To this end, the same procedures and experimental conditions described above were followed to prepare the lipid monolayers with an initial surface pressure of 15.0 ( 1.0 mN/m. An appropriate volume of bile salt solution (52.7 mol % NaTC/ 47.3 mol %NaGDC) was then injected into the subphase to increase the surface pressure to 18.0 ( 1.0 mN/m. The bile salt concentration in the subphase was ∼0.02 mM. After allowing bile salts to adsorb into the interface for 30 min and when the surface pressure had become essentially constant, a mixture of porcine pancreatic colipase and lipase at a molar ratio of 5:1 was carefully introduced into the subphase from the outside of the barriers. The concentration of colipase and lipase in the subphase was 50 and 10 nM, respectively. The monolayer was then equilibrated undisturbed for 30 min until the surface pressure was essentially constant before the monolayer was transferred onto a freshly cleaved mica sheet. Further additions of colipase-lipase solution to the subphase were made, and again the monolayer was equilibrated for 30 min before transferring it onto mica substrates. At high colipase and lipase concentrations (200 and 40 nM, respectively), the subphase was perfused with 1 L of 0.02 mM bile salt solution at a rate of ∼0.25 mL/s after allowing colipase and lipase to adsorb for 30 min. The perfusion was necessary because of the problem of the enzymes adsorbing directly onto the mica from the subphase during LB film transfer. In previous studies, at high protein concentrations, this passive adsorption was extensive enough to mask details on images of LB films.11 The bile salt solution for the perfusion was at a concentration identical to that of the subphase in order to avoid significant bile salt adsorption or desorption from the monolayer which would potentially interfere with the molecular organization at the interface. LB films were transferred when sufficient unadsorbed colipase and lipase had been removed from the subphase. AFM Imaging. AFM images were acquired in air at room temperature using a MFP-3D-BIO Asylum Research AFM (Asylum Research Ltd., Oxford, UK). Intermittent contact or tapping mode imaging was performed using etched silicon cantilevers (Olympus OMCL-AC160TS, Asylum Research Ltd., Oxford, UK) with a resonance frequency of ∼300 kHz, a spring constant of ∼42 N/m, and a tip radius of
