New insights into the pH-dependent interfacial adsorption of dog gastric lipase using the monolayer technique

Colloids and Surfaces B: Biointerfaces 111 (2013) 306–312

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New insights into the pH-dependent interfacial adsorption of dog gastric lipase using the monolayer technique Anaïs Bénarouche 1 , Vanessa Point 1 , Goetz Parsiegla, Frédéric Carrière, Jean-Franc¸ois Cavalier ∗ CNRS, Aix-Marseille Université, Enzymologie Interfaciale et Physiologie de la Lipolyse, UMR 7282, 31 chemin Joseph Aiguier, 13402 Marseille cedex 20, France

a r t i c l e

i n f o

Article history: Received 17 January 2013 Received in revised form 31 May 2013 Accepted 5 June 2013 Available online 20 June 2013 Keywords: Monomolecular film Enzyme adsorption Interfacial enzymology Kinetic parameters

a b s t r a c t The access to kinetic parameters of lipolytic enzyme adsorption onto lipids is essential for a better understanding of interfacial enzymology and lipase–lipid interactions. The interfacial adsorption of dog gastric lipase (DGL) was monitored as a function of pH and surface pressure (˘), independently from the catalytic activity, using non-hydrolysable 1,2-dilauroyl-sn-glycero-3-phosphocholine (DLPC) monomolecular films. The acid-stable DGL, which initiates fat digestion in the stomach, was then selected because its adsorption kinetics onto hydrophobic solid surfaces were already studied. This gastric lipase was therefore used as a model enzyme to validate both experimental and theoretical approaches. Results show that the adsorption process of DGL at the lipid/water interface depends on a pH-dependent adsorption equilibrium coefficient which is optimum at pH 5.0 (KAds = 1.7 ± 0.05 × 108 M−1 ). KAds values further allowed an indirect estimation of the molar fraction (˚E*(%) , mol%) as well as the molecular area (AE* ) of DGL adsorbed onto DLPC monolayer. Based on these data, a model for DGL adsorption onto DLPC monolayer at pH 5.0 is proposed for a surface pressure range of 15–25 mN m−1 . © 2013 Elsevier B.V. All rights reserved.

1. Introduction Lipases are water-soluble enzymes whereas their substrates are insoluble in water. The catalytic reaction of lipolysis is therefore related to various interfacial phenomena and depends strongly on the structural organization of the lipid substrates present at the interface, such as oil-in-water emulsions, membrane bilayers, micelles and vesicles [1–3]. Contrary to what is observed with “classical” enzymes acting on soluble substrates, the kinetic properties and substrate specificity of interfacial enzymes result from both the adsorption of the enzyme at the lipid/water interface and the interactions occurring between the substrate and the active site [1,4]. In other words, the rate of substrate hydrolysis by interfacial enzymes depends not only on the formation of an interfacial enzyme–substrate complex (E*S) and the catalytic constant (kcat ), but also on the adsorption (ka ) and desorption (kd ) rate constants of the enzyme to the lipid interface, as described in the kinetic model

Abbreviations: DGL, dog gastric lipase; DLPC, 1,2-dilauroyl-sn-glycero-3phosphocholine; HGL, human gastric lipase; TAG, triacylglycerol. ∗ Corresponding author. Tel.: +33 491164093; fax: +33 491715857. E-mail address: [email protected] (J.-F. Cavalier). 1 Both authors should be considered as equal first author, and are listed in alphabetical order. 0927-7765/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2013.06.025

developed by Verger and De Haas [5,6]. According to this interfacial model, variations of enzyme activity as a function of pH might result from changes in kcat and/or the enzyme–lipid interfacial adsorption equilibrium coefficient (KAds ). The understanding of the variation of lipolytic activities as a function of pH is a major challenge in view of the potential applications of lipases in the fields of chemistry [7,8], biotechnology [9,10] and medicine [11]. In the latter domain for instance, acidresistant lipases, such as gastric lipase, might be a good candidate in replacement enzyme therapy for the treatment of pancreatic enzyme insufficiency observed in chronic pancreatitis and cystic fibrosis [12–14]. Dog gastric lipase (DGL), which is active and stable at highly acidic pH 2.0 in the stomach environment where the gastrointestinal lipolysis of dietary fat is initiated, shows an optimum activity at pH 4.0 on long-chain triacylglycerols (TAGs) [15]. More recently, Chahinian et al. [16] showed that DGL can also act in aqueous solutions on short-chain vinyl butyrate with an optimum activity above pH 7.0, suggesting that this gastric lipase is able to hydrolyse ester bonds via the classical mechanism of serine hydrolases. Interestingly, in this latter study the optimum activity of DGL shifted towards lower pH values when vinyl butyrate concentration exceeded its solubility limit. Additional measurements on DGL adsorption onto solid hydrophobic surfaces using total internal reflection fluorescence (TIRF) and quartz crystal microbalance (QCM) confirmed the occurrence of a pH-dependent reversible

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adsorption process which was found to be optimum at pH 5.0 [16]. Taking into account these two independent kinetic studies on catalysis and adsorption on solid support, the authors concluded that the apparent optimum activity of gastric lipase at acidic pH reflected the adsorption process of the lipase at the lipid/water interface. Under physiological conditions, the first stage of lipid digestion is the formation of a fat emulsion which provides a higher specific surface for lipase adsorption. Dietary phospholipids, often present initially in lamellar structures, play an important role in the stabilization of fat emulsions by forming a monolayer at the surface of TAG droplets [17,18]. Such an interface is also commonly found in industrial food emulsions. The presence of phospholipids impairs the activity of pancreatic lipase on these substrates [19] but it has been clearly established that the release of fatty acids by gastric lipase promotes the action of pancreatic lipase on TAG droplets covered by phospholipids such as IntralipidTM (a soybean TAG emulsion stabilized with egg-yolk lecithins and used for parenteral intravenous infusions) or native milk fat globules [19,20]. Since the lipase adsorption at the lipid/water interface must occur before the insoluble substrate is hydrolysed, non-hydrolysable phospholipid monolayers spread at the air/water interface are very useful tools for studying such lipid–protein interactions independently from substrate hydrolysis [21–25]. The use of these model membranes allow to finely control several physical parameters such as pH of the subphase, lipid composition of the monolayer and density of lipids at the interface. The adsorption of a protein onto a phospholipid monolayer can be precisely monitored by following the increase in surface pressure (˘ max ) which is directly connected to the variation of lipid molecular area (i.e. lipid packing). By varying the initial surface pressure (˘ i ), it is therefore possible to analyse the influence of lipid packing on the interfacial binding of proteins. In view of the biochemical [15] and physiological properties [26–28] of DGL, and taking into account the preliminary results obtained on its adsorption on hydrophobic surfaces [16], DGL can thus be regarded as a model enzyme to study by the monolayer technique the influence of both pH and surface pressure on the kinetic parameters that regulate lipase adsorption at the lipid/water interface. We report here a simple and reliable method based on Langmuir adsorption isotherm [29] to monitor DGL binding onto a nonhydrolysable 1,2-dilauroyl-sn-glycero-3-phosphocholine (DLPC) phospholipid monolayer. This method allowed us not only to characterize the kinetic constants of lipase adsorption (ka , kd , KAds ), but also to estimate the related parameters (i.e., interfacial lipase concentration and molecular area) of the adsorbed enzyme. On the basis of these data, the cross-sectional areas of the adsorbed DGL molecule have been calculated, and a model for its binding to DLPC monomolecular film as a function of surface pressure is proposed.

2. Materials and methods 2.1. Materials 1,2-Dilauroyl-sn-glycero-3-phosphocholine (DLPC), from Avanti Polar Lipids Inc., was purchased from COGER (Paris, France) and was >99% purity. All other chemicals, including 2-(N-morpholino)ethane sulfonic acid (MES), Glycine, NaCl, CaCl2 , Tris, EDTA and sodium acetate, were purchased from Sigma–Fluka–Aldrich (St-Quentin-Fallavier, France) and were BioXtra grade. Chloroform (anhydrous for analysis, stabilized with amylene) was purchased from Carlo Erba Reactifs-SDS (Val de Reuil, France).

307

2.2. Lipase Crude recombinant dog gastric lipase (DGL) was provided by Meristem Therapeutics (Clermont-Ferrand, France) [30] and purified as described previously [31]. The concentration of purified DGL (4.094 mg mL−1 ) was determined by amino acids analysis performed at Laboratoire d’Enzymologie Moléculaire, Institut de Biologie Structurale (Grenoble, France). For the monolayer experiments, DGL stock solutions were prepared at a concentration of 1.1 mg mL−1 in 10 mM MES (pH 6.0) containing 150 mM NaCl. 2.3. Monomolecular film experiments 2.3.1. General methodology All experiments were performed at room temperature (25 ◦ C) using home-made Teflon troughs and the KSV5000 system (KSV, Helsinki, Finland) equipped with a temperature sensor probe, a Langmuir film balance and a mobile-barrier for compression isotherm experiments. The Wilhelmy method [32] was used to measure the difference of the surface tension existing between the surface of the aqueous phase ( 0 = 72.7 mN m−1 for pure water at 20 ◦ C) and that of the same aqueous phase covered by the phospholipid film (). This difference corresponds to the surface pressure: ˘ =  0 −  [33], directly monitored by the Langmuir film balance of the KSV5000 system. Surface pressure, barrier movement and temperature were monitored by the KSV Device Server Software v.1.20 installed on a computer running under Windows XP® . Before each experiment, the Teflon trough used was cleaned with tap water, and then gently brushed in the presence of distilled ethanol, before being washed again with tap water and abundantly rinsed with Milli-QTM water. Residual surface-active impurities were removed before each experiment by simultaneous sweeping and suction of the surface [34]. The aqueous subphase containing 100 mN NaCl, 21 mM CaCl2 and 1 mM EDTA was prepared with either 50 mM glycine–HCl for a pH value adjusted either at 2.0 or 3.0; 10 mM sodium acetate buffer for a pH value adjusted either at 4.0, 5.0 or 6.0; or 10 mM Tris buffer for a pH value of 7.0. The buffers were prepared with Milli-QTM water and filtered through a 0.45 ␮m Millipore membrane. The monolayer was prepared by spreading a few microliters of a DLPC solution (1 mg mL−1 in chloroform) until the desired initial surface pressure (˘ i ) was reached. The waiting time for the spreading solvent evaporation and for the film to reach equilibrium varied from 10 to 20 min depending on the spreading volume, the initial surface pressure, and the pH of the aqueous subphase. 2.3.2. Surface pressure-area isotherms The DLPC monolayer was formed over the surface of a homemade rectangular Teflon trough (volume 126.3 mL; surface area, 210.5 cm2 ) as described above. The surface pressure-area isotherms were recorded by monitoring the surface pressure increase during film compression with a mobile barrier at a constant rate of 10 mm min−1 (Fig. S1 in Supplementary data). Based on the trough dimensions and the amounts of lipids spread at the air/water interface, the barrier movement corresponded to a film compression of 3.5 A˚ 2 molecule−1 min−1 . The aqueous subphase was composed of the same buffers as described above. Each compression isotherm was performed in triplicate at each pH value, and the measurement of surface pressure was recorded every 5 s. 2.3.3. DGL adsorption onto DLPC monomolecular films The DLPC monolayer was formed over a home-made round Teflon trough (volume, 18.1 mL; surface area, 13.6 cm2 ) by spreading a DLPC chloroformic solution until the desired initial surface pressure (˘ i ) was reached (see Section 2.3.1). After injecting DGL into the aqueous subphase with an Hamilton syringe (from 5 ␮L

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to 80 ␮L DGL stock solution for final concentrations ranging from 5 nM to 90 nM, respectively), the surface pressure increase due to the adsorption of the lipase onto the DLPC monolayer was continuously recorded (every 5 s) until the equilibrium surface pressure was reached (around 60 min – Fig. S2 in Supplementary Data). The aqueous subphase was continuously stirred with a 1-cm magnetic bar rotating at 250 rpm. The system was designed so that the agitation of the aqueous phase with the magnetic bar did not disturb the stability of the surface film [35]. The aqueous subphase was composed of the same buffers as described above.

The molar fraction (˚E*(%) , mol%) of total lipase which is adsorbed onto the lipid monolayer can then be deduced from Eq. (6):

2.4. Treatment of kinetic data of DGL adsorption onto DLPC monolayer

AE∗ =

To analyse the DGL adsorption onto DLPC film, the experimental data (i.e. surface pressure increase with time – Fig. S2 in Supplementary Data) were fitted to the Langmuir equation for determining the rate constants of adsorption (ka , M−1 s−1 ) and desorption (kd , s−1 ). The Langmuir equation was adapted to ˘ measurements as follows (Eq. (1)):

ADLPC (Å2 molecule−1 ) is the molecular area of DLPC deduced from the surface pressure (˘)-molecular area isotherms at the final equilibrium surface pressure reached upon DGL adsorption; nDLPC is the number of DLPC molecules spread at the air/water interface at the working initial surface pressure (˘ i ); and  is the fractional surface coverage determined in Eq. (4). Eq. (7), thus obtained by means of our mathematical approach based on Verger’s kinetic model, therefore allowed assess to an estimation of the values of AE* (see Document S1 in Supplementary Data for the more details).

˘(t) = ˘i + ˘max ·



ka · CE ka · CE + kd



· [1 − exp (−(ka · CE + kd ) · t)](1)

where ˘ (t) is the surface pressure measured as a function of time; ˘ i is the initial surface pressure; ˘ max is the maximum variation of surface pressure reached upon DGL binding; and CE (mol L−1 ) is the enzyme concentration remaining in the subphase of the trough. In the monolayer system, since the ratio of surface area to volume of the trough (Strough /Vtrough ) is very small and close to 1 cm−1 , only a small fraction of the protein injected is expected to adsorb onto the lipid film [36–38]. Based on these findings, Verger et al. simplified their interfacial kinetic model [6,36,39] by making the assumption that CE remained almost unchanged and close to the concentration of the diluted enzyme after its injection into the subphase (CE0 ). With this approximation (CE ∼ = CE0 ), and using the Langmuir adsorption isotherm [29], Eq. (1) has been simplified to Eq. (2): ˘(t) = ˘i + ˘max ·  · [1 − exp (− · t)]

(2)

where  = ka · CE0 + kd

(3)

and =

ka · CE0 ka · CE0 + kd

(4)

( is the fraction of the total free adsorption (binding) sites coverage. Curve-fitting the experimental data points to Eq. (2), using KaleidaGraph 4.1 software from Synergy Software (Fig. S3A in Supplementary Data), allowed to calculate  values for multiple CE0 concentrations at each pH and ˘ i investigated. Values of ka and kd were then determined from the slope and y-intercept, respectively, of the linear plot of versus CE0 (Fig. S3B in Supplementary Data). The adsorption equilibrium coefficient, KAds , which represents the binding affinity between the protein and the lipid film, was obtained from the ratio of the measured rate constants (KAds = ka /kd = 1/KD ). Using the mathematical equations derived from the interfacial kinetic model of Verger et al. [39], it became possible to estimate, by indirect calculation, the theoretical value of the enzyme interfacial concentration ( E* , molecule cm−2 ) based on Eq. (5), where the trough volume (Vtrough ) is 18.1 mL and the surface area (Strough ) is 13.6 cm2 : E∗ =

CE0 (1 + KD ) · (Strough /Vtrough )

(5)

˚E∗(%) =

E∗ · (Strough /Vtrough ) CE0

=

100 1 + KD

(6)

Finally, the mathematical expression of the molecular area (AE* , Å2 molecule−1 ) occupied by the enzyme adsorbed onto DLPC films is: (Strough − ADLPC · nDLPC ) ·  E∗ · Strough

=

surface occupancy E∗

(7)

3. Results and discussion 3.1. Choice of DLPC as a model interface and surface pressure/area-isotherms of DLPC monolayer at various pH Gastric lipase is known to bind and hydrolyse lecithin-stabilized triacylglycerol emulsions such as IntralipidTM [16,40] but this lipase has no phospholipase activity. This enzyme is therefore able to penetrate the phospholipid layer surrounding lipid droplets in order to access to the triglyceride core. It is assumed that the interaction of gastric lipase with phospholipids is an important step in the overall process of interfacial catalysis. Phospholipid monomolecular films spread at the air/water interface can be used as biomimetic interfaces reproducing under controlled conditions the phospholipid monolayer surrounding various lipid bodies and emulsions. These films have been extensively used for studying lipase adsorption at non-hydrolysable interfaces, as well as the tensioactive properties of various proteins [21–25,41]. DLPC was selected in this study because it forms stable monolayers and has been often used for studying phospholipase activities using the monomolecular film technique [42–45]. Because DLPC is not a substrate for DGL, the enzyme binding at the lipid/water interface could be studied independently from the substrate hydrolysis step. As a prerequisite to study the pH-dependent adsorption of DGL to DLPC monolayers, the interfacial properties of this phospholipid at the air/water interface were analysed at pH values ranging from 2.0 to 7.0. Film stability was first checked by recording over 60 min the variations in surface pressure of a DLPC monolayer spread at various initial surface pressures, ˘ i . In all cases, no significant change in surface pressure (±0.2 mN m−1 ) was observed during the time course of the experiment whatever ˘ i or pH of the aqueous subphase applied. The surface pressure (˘)-molecular area (ADLPC ) curves deduced from compression isotherms were comparable or even superposable to those previously published [46–48] and were characteristic of a single liquid-expanded (LE) state at all surface pressures below the collapse (i.e. below 43 mN m−1 ) (Fig. S1 in Supplementary data). For ˘ i of 15, 20 and 25 mN m−1 used in adsorption kinetics, the enzyme will then bind DLPC in a single physical state, corresponding to the most favourable conditions to obtain accurate and reproducible ˘ measurements [41].

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Table 1 Kinetics constants derived from the adsorption kinetics of multiple DGL concentration onto DLPC films at pH 5.0 and at three initial surface pressures.a ˘ i (mN m−1 )

ka (M−1 s−1 )

kd (s−1 )

15 20 25

3.1 (±0.1) × 10 2.2 (±0.1) × 104 0.69 (±0.02) × 104 4

KAds (M−1 ) −4

1.2 (±0.04) × 10 1.2 (±0.03) × 10−4 2.2 (±0.05) × 10−4

2.5 (±0.1) × 108 1.7 (±0.1) × 108 0.32 (±0.01) × 108

a Values of the adsorption (ka ) and desorption (kd ) constants were determined from the slope and y-intercept, respectively, of the linear plot of  versus DGL concentration (Eq. (3) – Fig. S3 in Supplementary data). The adsorption equilibrium coefficient, KAds , which represents the binding affinity between the protein and the lipid film, was derived from the ratio of the measured rate constants (KAds = ka /kd ). Data are presented as mean values ± standard deviations (n = 2; CV% < 5.0%).

Fig. 1. Binding parameters of DGL into DLPC monomolecular film at pH 5.0. Variation of surface pressure increase (˘ max ) after DGL injection (20 nM final concentration) as a function of the initial surface pressure (˘ i ) of a DLPC monolayer. Data are presented as mean values ± standard deviations of two independent assays (n = 2; CV% < 5.0%).

3.2. Binding of DGL to DLPC monolayer Proteins binding capacity onto DLPC films can be precisely monitored following the increase in surface pressure (˘ max ) which is directly connected to the variation of DPLC molecular area (ADPLC , i.e. lipid packing). The influence of ˘ i on the adsorption/desorption process of DGL onto DLPC monolayers was studied at the optimal adsorption pH value 5.0, as reported by Chahinian et al. [16]. The optimal final protein concentration used in these experiments, was determined by performing ˘ measurements at different lipase concentrations which allowed to obtain a DGL surface saturation of 20 nM (Fig. S5A in Supplementary Data). The plot ˘ max = f(˘ i ) depicted in Fig. 1, was first used to evaluate the binding parameters of DGL. Linear extrapolation to zero surface pressure increase (˘ max = 0) allowed to determine the critical surface pressure (˘ c ) [35], also named “maximum insertion pressure” [41], as being equal to 30.3 mN m−1 . Above this ˘ c value, which is specifically related to the protein and the lipid forming the monolayer, no increase in the surface pressure will then occur [35]. The synergy (slope of the linear regression +1) and ˘ 0 (y-intercept of the curve) as defined by Salesse et al. [41,49,50] can also be deduced from the same plot. The observed positive synergy value (0.22) and the occurrence of a ˘ 0 value (23.7 mN m−1 ) lower than the related ˘ c (30.3 mN m−1 ) is consistent with an ‘insertion’ surface pressure and favourable interactions between the protein and the phospholipid film. Consequently DGL is able to bind into DLPC monolayers at high surface pressures and is not excluded from the film. In previous reports [21,38] native human gastric lipase (HGL) was shown to adsorb onto monomolecular film of long chain egg PC with a

critical surface pressure ˘ c of around 22.0 mN m−1 [35,51] but was not studied using DLPC films. We have shown in additional experiments that DGL exhibits a similar ˘ c value of 21.5 mN m−1 onto Egg PC monolayers at pH 5.0 (Fig. S4 in Supplementary Data). These results suggest that both DGL and HGL may have a similar behaviour in presence of phospholipid films. However, the differences in the ˘ c values of DGL obtained with egg PC (21.5 mN m−1 ) and DLPC (30.3 mN m−1 ) could result from the chemical structure of acylchains in each phospholipid: the longer and unsaturated egg PC chains would reduce DGL adsorption onto the PC monolayer. 3.3. Determination of the adsorption kinetic parameters of DGL onto DLPC monolayer Taking into account the latter determined ˘ c values, the use of DLPC monolayers gave access to the kinetic parameters of DGL adsorption at surface pressures above 21 mN m−1 , which would be impossible to obtain with egg PC films. In order to investigate the binding process of this gastric lipase as a function of the pH of the subphase, a series of adsorption kinetics were performed at ˘ i = 20 mN m−1 with varying DGL concentrations in the range of 5.0 to 90.0 nM. Kinetic adsorption (ka ) and desorption (kd ) rate constants were determined for each pH by curve-fitting experimental data points to the Langmuir adsorption equation (Eq. (2)). Both constants were maximal at pH 2.0 and decreased significantly with increasing pH values (Fig. 2A and B). Minimal values for adsorption (ka = 2.16 × 104 M−1 s−1 ) and desorption (kd = 1.24 × 10−4 s−1 ) rate constants were obtained at pH 5.0, where the adsorption equilibrium coefficient, KAds (1.7 × 108 M−1 ; Fig. 2C) reached its maximum. In particular, it should be noticed that this optimum KAds value obtained from our adsorption kinetics at pH 5.0 and ˘ i = 20 mN m−1 , is nearly identical to the one (1.5 × 108 M−1 ) previously found by Chahinian et al. [16]. The adsorption kinetic constants were further measured at ˘ i of 15 and 25 mN m−1 (Table 1 and Fig. S3B in Supplementary Data). The maximum desorption rate constant (kd = 2.2 × 10−4 s−1 ) was obtained at 25 mN m−1 , whereas no significant variations in kd values occurred

Table 2 Values related to DGL adsorption onto DLPC films at various pH.a pH

˘ i (mN m−1 )

DGL adsorption ˘ max (mN m−1 )

2 3 4 5 5 5 6 7

20 20 20 15 20 25 20 20

7.9 8.6 8.7 12.6 8.2 3.6 7.6 8.4

± ± ± ± ± ± ± ±

0.1 0.3 0.05 0.5 0.2 0.03 0.2 0.3

 E* (molecule cm−2 )

AE* (Å2 molecule−1 )

1.39 (±0.04) × 10 2.01 (±0.07) × 1012 1.51 (±0.06) × 1012 3.23 (±0.09) × 1012 2.37 (±0.06) × 1012 0.49 (±0.01) × 1012 0.81 (±0.03) × 1012 0.95 (±0.05) × 1012

1084 494 771 793 718 674 1395 1498

12

± ± ± ± ± ± ± ±

30 16 32 22 19 12 61 82

˚E*(%) (mol%) 8.7 12.6 9.4 20.2 14.8 3.1 5.1 5.9

± ± ± ± ± ± ± ±

0.2 0.4 0.4 0.9 1.2 0.2 0.2 0.3

a The maximum surface pressure increase (˘ max ) was deduced from the respective experimental adsorption curves.  E* , ˚E*(%) and AE* values were estimated by indirect calculation using Eqs. (5)–(7), respectively, for a DGL final concentration (CE0 ) of 20 nM. Data are expressed as mean values ± standard deviations (n = 2; CV% < 5.0%).

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Fig. 3. Variations of the calculated molecular areas (AE* ) of the adsorbed DGL onto DLPC films (A) as a function of pH at ˘ i = 20 mN m−1 , and (B) as a function of ˘ i at pH 5.0. AE* values were estimated by indirect calculation using Eq. (7) for a DGL final concentration (CE0 ) of 20 nM. Data are presented as mean values ± standard deviations of two independent assays (n = 2; CV < 5.0%).

plateau value corresponding to the saturation of the monolayer interface with the protein (Fig. S5B in Supplementary Data). 3.4. Estimation of the amount of DGL adsorbed onto DLPC monolayers

Fig. 2. Effects of pH on the kinetic and association constants of DGL binding onto DLPC monolayer at an initial surface pressure of 20 mN m−1 . The adsorption constant (ka , panel A), the desorption constant (kd , panel B) were obtained by curve-fitting experimental data points to the Langmuir adsorption equation (Eq. (2)). The adsorption equilibrium coefficient (KAds , panel C) was calculated from the ratio of both rate constants. DGL final concentration in the subphase ranged from 5.0 to 90.0 nM. Data are presented as mean values ± standard deviations of two independent assays (n = 2; CV% < 5.0%).

between 15 and 20 mN m−1 (mean kd ∼ = 1.2 × 10−4 s−1 ). In contrast, ka values decreased from 15 mN m−1 (ka = 3.1 × 104 M−1 s−1 ) to 25 mN m−1 (ka = 0.69 × 104 M−1 s−1 ) (Table 1). The resulting adsorption equilibrium coefficient KAds was found to follow a similar decreasing dependency upon the initial surface pressure from 15 to 25 mN m−1 (Table 1). From these latter experiments we could determine the binding capacity (˘/t)t = 0 of the protein as the initial rate of surface pressure increase after protein injection [35,52,53]. For each of the three ˘ i tested, (˘/t)t = 0 was directly correlated with the amount of lipase in the subphase. Above the threshold value of 20 nM protein concentration, the (˘/t)t = 0 rates reached a

The experimentally-determined kinetic parameters (ka , kd , KAds ) made it also possible to estimate the theoretical DGL interfacial concentration ( E* , molecule cm−2 ) and to deduce the molar fraction of adsorbed lipase (˚E*(%) , mol%) onto DLPC films (Table 2), using Eqs. (5) and (6), respectively, derived from the interfacial kinetic model developed by Verger et al. [5]. As expected,  E* and ˚E*(%) values have the same pH and surface pressure dependencies as KAds . When calculating ˚E*(%) at ˘ i = 20 mN m−1 in a pH range from 2.0 to 7.0, the maximum amount of absorbed DGL (˚E*(%) = 14.8 ± 1.2 mol%) was reached at pH 5.0, and the lowest one (˚E*(%) = 5.1 ± 0.2 mol%) at pH 6.0 (Table 2). At pH 5.0, DGL adsorption was found to decrease with increasing surface pressures. More precisely, ˚E*(%) dropped from 20.2 ± 0.9 mol% at 15 mN m−1 to 3.1 ± 0.2 mol% at 25 mN m−1 (Table 2). The surface areas occupied by a single molecule of the adsorbed protein (AE* ) were further calculated using Eq. (7) and were then found to vary significantly with pH (Fig. 3A and Table 2). At pH 4–5, where DGL exhibited its highest lipolytic activity [15], a medium range AE* value of 744 ± 26 A˚ 2 molecule−1 was obtained. At pH 5.0, AE* values decrease almost linearly from 793 to 674 A˚ 2 molecule−1 within the range 15–25 mN m−1 (Fig. 3B and Table 2). These decreasing curve patterns of both  E* and subsequently ˚E*(%) values (Table 2), as well as AE* values at pH 5.0 in the ˘ i -range of 15–25 mN m−1 are in agreement with the general behaviour of lipolytic enzymes showing a decrease in their binding

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Fig. 4. Putative orientation of DGL adsorbed onto a DLPC monolayer spread at the air/water interface. The structure of the DGL is shown in molecular surface representation. Hydrophobic residues (Ala, Leu, Ile, Val, Trp, Tyr, Phe, Pro, Met) are highlighted in white, and the catalytic Ser153 is coloured in red. (A) Side view of the DGL molecule oriented at the lipid interface. (B) Top view of the DGL showing the hydrophobic ring surrounding the active site entrance and parallel to the putative lipid interface. (C) Top views of DGL with cross-sections (grey colour) made parallel to the interface plane. The position of cross-sections and their distances from the top of the DGL molecule are indicated in panel A. In panel C, the cross-section areas are associated with the surface pressures for which similar values of molecular areas (AE* ) were estimated using Eq. (7). These models were drawn using PyMOL Molecular Graphics System (Version 1.3, Schrödinger, LLC) and the 1K8Q Protein Data Bank file for DGL. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of the article.)

capacity at high surface pressure (i.e., more compact lipid packing) [6,34,38,54]. All these findings therefore validate our experimental and theoretical approach to quantify the adsorption of lipases onto lipid monolayers. In particular, it is noteworthy that the observed amount of DGL (˚E*(%) = 3.1%) bound to DLPC film at 25 mN m−1 was nearly identical to the one previously determined by ELISA tests with HGL (˚E*(%) ∼ = 2.8%) on 1,2-dicaprin (i.e. lipid substrate) films under the same operating conditions [38]. 3.5. Model of DGL adsorption onto a DLPC monolayer The DGL 3D structure (PDB ID: 1K8Q) was previously obtained in its active conformation with an open lid [31,55]. This change in the conformation of the lid, that controls access to the active site, was established by comparison with the 3D structure of HGL [56,57] in which the lid is found to be closed. The opening of this lid is achieved by a 180◦ rotation of helices H1 (residues 217–228) and H2 (residues 231–244) around the axis of H2 helix. As a result, the polar face of the two helices is then buried, whereas the hydrophobic part becomes exposed, as previously described in the case of other lipases [1]. In consequence, a large hydrophobic patch (nearly 2227 A˚ 2 ) surrounding the entrance of the active site becomes exposed at the surface [31,55]. When this open 3D structure of DGL was aligned with that of the HPL-colipase complex, based on the ␣/␤ hydrolase fold and the catalytic triad, the hydrophobic ring surrounding the entrance to the DGL active site was found to be in the same plane as the interfacial recognition site (IRS) of HPL, including the open lid domain, the ␤9 loop and the C-terminal domain ␤5 loop [1]. This also allowed to speculate on DGL orientation at the lipid/water interface since a model

for HPL orientation was previously proposed [58]. This orientation of HPL was based on structural homology between HPL C-terminal domain and the C2 domain of cPLA2 [58], and the orientation of this C2 domain bound to DOPM membrane determined by side-directed spin-labelling and EPR spectroscopy [59]. From this superimposition, it was found that the hydrophobic loops of HPL, potentially involved in IRS came in close contact with the interfacial plane. The orientation of open DGL deduced from these previous observations therefore appeared as a good model for illustrating DGL adsorption onto a DLPC monolayer. The DGL molecule was first oriented with its hydrophobic ring parallel to the putative lipid/water interface (Fig. 4A and B), and stepwise cross-sections distant by 1.5 A˚ were made parallel to this plane (Fig. 4C). The cross-section areas were then compared to the AE* values estimated from DGL adsorption onto DLPC monolayers and an interesting similarity was observed. Indeed, the areas of the cross-sections made at 10.0, 11.5 and 13.0 A˚ 2 from the top of the DGL molecule (Fig. 4A) had the same values as the molecular areas (AE* ) extrapolated from DGL adsorption at 25, 20 and 15 mN m−1 , respectively (Fig. 3B). Although it remains speculative, the anti-correlation between surface pressure and the calculated distance of DGL cross-section from the top of the molecule might reflect a decreasing binding capacity of the lipase when the surface pressure is increased. Several peptide stretches part of the hydrophobic ring surrounding the active site entrance and potentially involved in the IRS of DGL (i.e.: Leu1 to Lys4 N-terminal residues; Phe217-Ser228 including ␣e3 helix from the open lid; Phe201-Phe212 including residues from ␣e2 helix [31,57,60]) are found above the DGL cross-sections (Fig. 4), suggesting they would be always in contact with the lipid interface.

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4. Conclusion All these findings corroborate that lipase adsorption is highly dependent on the “interfacial quality” of the lipid/water interface [61] which is mostly related to the orientation and conformation of the lipids forming the monolayer as well as their molecular packing linked to surface pressure. The present data show that DGL adsorption onto a phospholipid monolayer depends on a pHdependent adsorption equilibrium coefficient, KAds . Interestingly, KAds values obtained here by tensiometry measurements are similar to those previously reported in the course of DGL adsorption onto hydrophobic surfaces and determined by using surface spectroscopy (TIRF) [16]. It therefore seems that the pH-dependence adsorption of gastric lipase is optimum at acidic pH whatever the interface. This will reinforce the assumption that the optimum activity of gastric lipase at low pH mainly results from the lipase adsorption step at the lipid/water interface. Determining adsorption constants with a real lipase substrate, such as 1,2-dicaprin monolayer, remains a challenge. However, previous data obtained from the recovery of 1,2-dicaprin monolayer in the presence of HGL suggest that gastric lipase might bind similarly to 1,2-dicaprin and DLPC monolayer, thus supporting the use of phospholipid monolayers as model interfaces for studying gastric lipase adsorption step. More generally, the evaluation of kinetic parameters of enzyme adsorption onto a lipid monolayer, now fully accessible with our developed method, brings new insights in the understanding of lipase mechanism of action and should open new prospects in the investigation of lipase–lipid interactions. Acknowledgements A. Bénarouche was supported by a PhD fellowship from the Ministère de l’Enseignement Supérieur et de la Recherche. This work was supported by the CNRS and by the LISA Carnot Institute (Convention ANR n◦ 07-CARN-009-01). The authors also wish to thank Dr. Jessica Blanc for revising the English manuscript. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb. 2013.06.025. References [1] A. Aloulou, J.A. Rodriguez, S. Fernandez, D. van Oosterhout, D. Puccinelli, F. Carrière, Biochim. Biophys. Acta 1761 (2006) 995–1013. [2] H. Singh, A. Ye, D. Horne, Prog. Lipid Res. 48 (2009) 92–100. [3] M. Golding, T.J. Wooster, Curr. Opin. Colloid Interface Sci. 15 (2010) 90–101. [4] P. Reis, K. Holmberg, H. Watzke, M.E. Leser, R. Miller, Adv. Colloid Interface Sci. 147/148 (2009) 237–250. [5] R. Verger, G.H. de Haas, Chem. Phys. Lipids 10 (1973) 127–136. [6] R. Verger, M.C.E. Mieras, G.H. de Haas, J. Biol. Chem. 248 (1973) 4023–4034. [7] K. Faber, Bio-transformations in Organic Chemistry, 4th ed., Springer-Verlag, Berlin-Heidelberg, 1992. [8] R.D. Schmid, R. Verger, Angew. Chem. Int. Ed. 37 (1998) 1608–1633. [9] E. Vulfson, Industrial applications of lipases, in: Lipases: Their Structure, Biochemistry and Application, Cambridge University Press, Cambridge, UK, 1994, pp. p271–p288. [10] F.X. Malcata, Engineering of/with Lipases, Kluwer Academic Publishers, Dordrecht, 1995. [11] H. Lengsfeld, G. Beaumier-Gallon, H. Chahinian, A. De Caro, R. Verger, R. Laugier, F. Carrière, Physiology of gastrointestinal lipolysis and therapeutical use of lipases and digestive lipase inhibitors, in: G. Müller, S. Petry (Eds.), Lipases and Phospholipases in Drug Development, Wiley-VCH, Weinheim, 2004, pp. 195–223. [12] M. Roulet, A.M. Weber, Y. Paradis, C.C. Roy, L. Chartraud, R. Lasalle, C.L. Morin, Pediatr. Res. 14 (1980) 1360–1362. [13] F. Carrière, P. Grandval, C. Renou, A. Palomba, F. Priéri, J. Giallo, F. Henniges, S. Sander-Struckmeier, R. Laugier, Clin. Gastroenterol. Hepatol. 3 (2005) 28–38.

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