Journal of Colloid and Interface Science 488 (2017) 149–154 Contents lists available at ScienceDirect Journal of Colloid and Interface Science journal homepage: www.elsevier.com/locate/jcis Regular Article A comparative study on kinetics and substrate specificities of Phospholipase A1 with Thermomyces lanuginosus lipase Ruipu Xin a, Faez Iqbal Khan b, Zexin Zhao c, Zedong Zhang a, Bo Yang c, Yonghua Wang a,⇑ a College of Food Sciences and Engineering, South China University of Technology, Guangzhou 510640, PR China School of Chemistry and Chemical Engineering, Henan University of Technology, Zhengzhou 450001, PR China c School of Bioscience and Bioengineering, South China University of Technology, Guangzhou 510006, PR China b g r a p h i c a l a b s t r a c t DOPC Air h Water √ C-terminal Region TLL PLA1 C-terminal region played significant role in enzyme acvity and substrate specificity. a r t i c l e i n f o Article history: Received 4 August 2016 Revised 20 October 2016 Accepted 20 October 2016 Available online 21 October 2016 Keywords: Lipase Phospholipase A1 Chain-length specificity Regiospecificity Monolayer Interfacial binding Substrate specificity a b s t r a c t The mechanism of lipase binding to the lipid-water interface is crucial for substrate specificity and kinetic properties. In this study, the chain-length specificity, regiospecificity and substrate specificity of Phospholipase A1 (PLA1) and its parent enzyme Thermomyces lanuginosus lipase (TLL) have been investigated using a classical emulsion system. The results show that both PLA1 and TLL are 1,3-regioselective lipases. Additionally, the hydrolytic activity of PLA1 is comparatively lower on short-chain triacylglyceride (TAG) and higher on phosphatidylcholine (PC) than the hydrolytic activity of TLL. Further, the results obtained with monolayer film techniques demonstrate that the C-terminal region regulates the binding of PLA1 to PC. A hypothesis is presented according to which the a9 helix of C-terminal region in PLA1 not only controls the opening of lid but also serves as a membrane anchor that assists in binding to PC. These findings bring new insight into rational design of novel lipases with intriguing functionalities. Ó 2016 Elsevier Inc. All rights reserved. Abbreviations: PLA1, phospholipase A1; TLL, Thermomyces lanuginosus lipase; TAG, triacylglyceride; PC, phosphatidylcholine; FOL, Fusarium oxysporum lipase; RML, Rhizomucor miehei lipase; TC4, tributyrin; TC10, trioctanoin; TC18, triolein; DOPC, 1,2-dioleoyl-sn-glycero-3-phosphocholine; b-CD, b-cyclodextrin; YLLIP8, Yarrowia lipolytica LIP8; CBD, cellulose binding domain; GZEL, Gibberella zeae lipase; kcat/KM, catalytic efficiency; KM, substrate affinity constant; FA, fatty acid; LCFA, long chain fatty acid. ⇑ Corresponding author. E-mail address: yonghw@scut.edu.cn (Y. Wang). http://dx.doi.org/10.1016/j.jcis.2016.10.058 0021-9797/Ó 2016 Elsevier Inc. All rights reserved. 150 R. Xin et al. / Journal of Colloid and Interface Science 488 (2017) 149–154 1. Introduction Lipases (E.C. 3.1.1.3) are interfacial enzymes that catalyze the hydrolysis of lipids at the lipid-water interface [1,2,3]. The reaction involves binding of the lipase to the lipid-water interface [4], which is crucial in controlling substrate specificity and activity by altering both, the structure of the lipase and the physical state of the interface [5]. Compared with enzyme reactions occurring in homogeneous phase, lipase reactions are fairly difficult to investigate. Various effects can interfere with the hydrolytic reaction such as the organization of the substrates in emulsions, monolayers, micelles, vesicles or liposomes [3]. Mostly, two approaches have been used to measure the kinetic properties of lipases, i.e. the pH-stat method using emulsified substrates and the baro-stat with lipid films (monolayer film technique). Using the pH-stat method in emulsion systems, the migration of lipases between lipid particles, substrate replenishment, alteration and destabilization of lipid particles affect the kinetic properties of lipases [6]. Monolayer techniques enable control of interfacial properties, such as the orientation and packing of the molecules, the changes in lateral density as well as the variations in lipid organization and structure [7,8], which also influence the substrate specificity of lipases [8–12]. Therefore, both methods provide complementary information on the kinetic properties of lipases. The action of lipase binding to the lipid-water interface has been studied extensively, revealing a prominent role of the C-terminal domain also influences substrate specificity and activity of lipases [4,5,13–15]. Phospholipase A1 (PLA1), which is a fusion protein of Thermomyces lanuginosus lipase (TLL) and an extra C-terminal region of Fusarium oxysporum lipase (FOL) [16], has been described to undergo interfacial activation [17]. Despite of the importance of both PLA1 and TLL in various industrial processes [18–27], a comparative study of the kinetic properties of both lipases is still missing today. Therefore in this study, the amino acid sequence and structure of PLA1 and TLL were analyzed. Particularly, the different substrate scopes of PLA1 and TLL were investigated using the classical emulsion system. Furthermore, the interfacial properties of PLA1 and TLL were investigated on TAG and PC films using monolayer film technique. 2. Material and methods 2.1. Chemicals Soybean-phosphatidylcholine (Soybean-PC) was purchased from Avanti Polar Lipids (Alabama Alabaster, America). Tributyrin (TC4, 99%), trioctanoin (TC10, 99%), triolein (TC18, 99%), 1,2dioleoyl-sn-glycero-3-phosphocholine (DOPC, 99%), chloroform (99%) and methanol (99%) were purchased from Sigma-Aldrich (Shanghai, China). Soybean oil was obtained from Donlinks (Guangzhou, China). b-cyclodextrin (b-CD) and microcrystalline cellulose were purchased from Aladdin (Shanghai, China). NaOH and ethanol were procured from Guangzhou Chemical Reagent Factory (Guangzhou, China). Milli-Q water (P18.2 MX cm) was used in all experiments. All other chemicals were of analytical grade. 2.2. Lipases purification The gene of PLA1 was previously synthesized by Sangon (Shanghai, China) [28]. The PLA1 lipase gene was cloned downstream of the cellulose binding domain (CBD) tag in the pET23a vector as described previously [29]. PLA1 with CBD tag (PLA1-CBD) was expressed in E. coli strain SHuffleÒ T7 expression system. The purification of PLA1 was performed as described previously by Lan et al. [29]. Accordingly, crude enzymes of T. lanuginosus lipase (TLL, Lipozyme TL100L) were provided by Novozyme (Copenhagen, Denmark). TLL was purified using a Q Sepharose column (GE, Boston, USA) as described by Qin et al. [30]. SDS-PAGE analysis of the purified PLA1 and TLL proteins showed a predominant protein band with the apparent molecular weights of approximately 35 and 30 kDa, respectively (Fig. S1). The enzyme solutions of PLA1 and TLL were concentrated by ultrafiltration and exchanged with 0.1 M PBS buffer (pH 7.0). Protein concentrations were determined by the Bradford method using bovine serum albumin as a reference [31]. 2.3. Sequence and structure analysis The amino acid sequences of related lipases were aligned using the Blast search and alignment tool of the Universal Protein Knowledge Base (www.uniprot.org), and presented using ESPript [32,33]. The protein structures of PLA1 and TLL were aligned and visualized using PyMol (DeLano Scientific LLC) [34]. 2.4. Lipase activity measurements Lipase activity was assayed with a pH-stat device (Radiometer, Copenhagen, Denmark) in a thermostatic vessel (25 °C). Each assay was performed with a mechanically stirred emulsion of TAG or PC as substrate. The specific activity of lipase was determined by titrating the free fatty acids (FAs) liberated during the hydrolysis reaction by PLA1 and TLL. The specific activity was expressed in units (U) per milligram of lipase. One U corresponds to 1 lmol of free fatty acid released per minute. The values are presented in mean ± standard deviation based on three independent experiments. 2.5. Kinetic measurements by monolayer film technique Micro TroughX Langmuir-Blodgett trough procured from Kibron Inc. (Espoo, Finland) was used to measure the enzymatic kinetics at 25 °C. The Teflon trough is composed of a reaction compartment (volume, 3 mL; surface area, 12.56 cm2) and two reservoir compartments (surface area, 55.46 cm2), both of which are connected with the reaction compartment by one narrow surface channel. A force probe comprised of a tensometric cantilever was placed the interface in the center of the trough to measure the surface pressure based on Wilhelmy plate method [9]. The precision in the measurement of the surface pressure by the Wilhelmy plate method was ±0.1 mN m 1. The trough and barriers were rinsed with ethanol and Milli-Q water prior to use. The surface pressure of the interface was adjusted to zero. Residual surface-active impurities on the Teflon trough were removed before each experiment by simultaneously sweeping and suction of surface [9]. Experiments proceeded after the change in surface pressure was less than 0.5 mN m 1. A magnetic stirrer (0.5-cm) was used to stir the subphase at 250 rpm. The aqueous subphase contained PBS buffer (10 mM, pH 7.0), which was prepared with Milli-Q water and filtered through a 0.45-lm Millipore membrane. The DOPC and triolein were dissolved quickly in chloroform/methanol solution (3:1 v/v) at 20 °C, as described previously [35–37]. The monolayer was prepared by spreading 20 lL of DOPC solution (0.2 g L 1) using a Hamilton microsyringe. The waiting time for the solvent evaporation varied between 10 and 15 min. The desired initial surface pressure (pi) was reached by moving two barriers at a rate of 10 mm min 1. The enzyme solution was injected into the subphase R. Xin et al. / Journal of Colloid and Interface Science 488 (2017) 149–154 after organic solvents were evaporated. Enzymatic activities were determined using the baro-stat technique on a ‘‘zero-order” trough [38]. The surface activity was defined as the amount of substrate that was hydrolyzes per minute, per surface area in the reaction compartment, which contained 1 M of lipase. All experiments were carried out in triplicate. 3. Results and discussion 151 activation mechanism [40]. Because the amino acid residue Asp294 of the C-terminal helix and the Arg87 of the lid a3 helix are conserved in PLA1 and GZEL (Fig. 1B), a similar ‘double-lock’ activation mechanism may be hypothesized for PLA1 as well. Lipases without the extra C-terminal helix (TLL and RML) should follow the normal activation mechanism. Notably, the C-terminal helix (Asp294) of PLA1 may transiently interact with the lid a3 helix (Arg87) and play a critical role in controlling the lid opening and enzyme activity. 3.1. Sequence alignment and structural comparison 3.2. Chain-length specificity and regiospecificity on TAG PLA1 has a high amino acid similarity with Gibberella zeae lipase (GZEL) (94.14%), TLL (84.38%) and RML (56.92%), respectively. The three-dimensional structure of PLA1 has been modeled using the crystal structures of the above mentioned lipases as templates by An et al. [18]. The structure of PLA1 was superimposed on TLL with a RMSD for Ca of 0.397 Å. One of the key differences observed between the two lipases is that PLA1 possesses an additional C-terminal region composed of a 55-residue extension (Asn269-Ala315) (Fig. 1). The flexible coil region (Asn269-Thr291) of GZEL [39,40] appears to be partially disordered in PLA1. The coil sequence followed by the helical region (Asp292-Lys311) of PLA1 is expected to temporarily interact with the enzyme surface, as observed with other lipases [15,41]. Similar C-terminal extensions have also been reported in other lipases [15]. In the closed form of GZEL, the a9 helix (Asp293) interacts with the a3 helix (Arg86) [40]. Lou et al. proposed a novel ‘double-lock’ The regioselectivity of both PLA1 and TLL on TAGs was as expected [42–45]. During the hydrolysis of soybean oil no evidence for 1,3-DAG was found confirming the reported 1,3-selectivity of the enzymes. To investigate the chain-length specificity of PLA1 and TLL, three different TAGs (TC4, TC10 and TC18) were selected. PLA1 was mainly active on medium-chain TAGs (TC10) and less active on short-chain TAGs (TC4) (Table 1 and Fig. 2). The specific activities of PLA1 were 477.04, 3092.35 and 1463.11 U/mg with TC4, TC10 and TC18 as substrates, respectively. However, TLL hydrolyses more efficiently the short-chain TAGs. One possible explanation is that PLA1 presents a structure that inefficiently binds to short-chain TAGs. The ‘lockhole-hinge-lock pin’ activity switch in PLA1 may be locked by an accessory C-terminal a-helix, therefore decreasing the penetration capacity into the interface of TC4 to activate the lipase [9]. Therefore, we Fig. 1. (A) Overlay of PLA1 with the C-terminal helical extension (red) and TLL (green) structures represented as cartoon form. (B) Sequence alignment of PLA1 and TLL (PDB id:1EIN). Secondary structural elements of the lipase crystal structure are shown at the top of the alignment. Arrows indicate b-strands, and helical curves denote a-helices. Residues highlighted in red background are identical among the compared proteins, residues highlighted in red are conserved, and the lock residues are highlighted by green frames, respectively. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.) 152 R. Xin et al. / Journal of Colloid and Interface Science 488 (2017) 149–154 Table 1 Chain length selectivity of PLA1 and TLL by emulsified method. Lipase Specific activitya TC4 TC10 TC18 PLA1 TLL 477.04 ± 20.43 5105.05 ± 89.26 3092.35 ± 49.24 7702.71 ± 30.22 1463.11 ± 80.91 4091.32 ± 120.34 TC4/TC18 ratio TC10/TC18 ratio 0.33 1.25 2.11 1.88 a Specific activity was determined by emulsified method and expressed in units (U) per milligram of lipase. One U corresponds to 1 lmol of free fatty acid released per minute. The values were mean ± standard deviation based on three independent experiments. Fig. 2. Chain-length specificity of PLA1 and TLL. Activity measurements were performed using TC4, TC10 or TC18 emulsion. assume that the C-terminal region of PLA1 may be involved in interfacial activation and affect the chain-length specificity. 3.3. Substrate specificities on PC/TAG The hydrolytic activities of PLA1 and TLL were measured using PC (soybean-PC) and TAG (soybean oil) emulsions at pH 7.0 and 25 °C. PLA1 was found to display both higher phospholipase activity (4403.00 ± 30.24 U/mg) and lipase activity (8728.31 ± 300.71 U/mg) than TLL. In addition, PLA1 displayed a higher ratio of phospholipase to lipase activity (0.504) compared with TLL (0.00685) (Table 2). 3.4. Kinetic studies using the emulsion method To explain the different substrate specificities towards PC/TAG between PLA1 and TLL, the kinetic parameters of lipase were determined using Lineweaver-Burk plots (Table 3). The catalytic efficiency (kcat/KM) of PLA1 and TLL were found to be similar with TAG as substrate. However, the kcat/KM of TLL was 6.08-fold lower than that of PLA1 using PC as substrate. Furthermore, the substrate affinity constant (KM) of TLL on PC was 3.67-fold lower compared with that of PLA1. Table 2 Substrate specificities on PC/TAG of PLA1 and TLL. Lipase PLA1 TLL Specific activitya PC/TAG ratio PC TAG 4403.00 ± 30.24 136.5100 ± 14.26 8728.3100 ± 300.71 19930.8300 ± 500.23 0.504 0.00685 a Specific activity was determined by emulsified method and expressed in units (U) per milligram of lipase. One U corresponds to 1 lmol of free fatty acid released per minute. 3.5. Kinetic measurements by monolayer film technique The monolayer technique is an interfacial tensiometry method, simultaneously monitoring and controlling the surface pressure. Hence, this method also allows to influence physicochemical parameters of the substrate such as molecular orientation and molecular density [45]. Therefore, determination of the interfacial properties of PLA1 and TLL by the monolayer technique can be helpful to further explain the different substrate specificities towards PC/TAG. PLA1 and TLL lead to the conversion of TAG into Sn-2 monoglyceride (2-MAG) and FA. Hence, the reaction mixture is composed of at least four surface-active ingredients (TAG, MAG, FA and lipase) leading to a complex equilibrium [46]. A competition for the interface among lipases, long chain fatty acids (LCFAs) and 2-MAGs occurs. Previous results have demonstrated that 2-MAG excludes both TAG and lipase from the interface [47]. It is worth mentioning here that in these experiments a low concentration of b-CD (0.7 mM) was applied to the aqueous subphase to facilitate the solubilization of MAG and LCFA [48]. Karray et al. have shown that the b-CD in the aqueous phase has not effect on the stability of the monolayer [8,47]. PLA1 and TLL on a triolein monolayer showed a maximum activity of 1.49 and 1.50 mol cm 2 M 1 min 1 at the a surface pressure of 25 and 20 mN m 1, respectively (Fig. 3A). PLA1 showed a bell shaped curve on the DOPC substrate with a characteristic optimal surface pressure (Fig. 3B). The highest hydrolytic activity of PLA1 and TLL on DOPC monolayer was 0.40 and 0.017 mol cm 2 M 1 min 1, respectively. The initial increase in hydrolytic activity at surface pressure may be ascribed to an improvement of enzyme-substrate binding, whereas the decrease of hydrolysis at high surface pressure may due to poor penetration capacity of the lipase into the surface [49,50]. Furthermore, the hydrolytic activity of TLL measured on the DOPC films was very low even at high lipase loadings. Furthermore, a 153 R. Xin et al. / Journal of Colloid and Interface Science 488 (2017) 149–154 Table 3 Kinetic parameters of PLA1 and TLL by emulsified method. a b c d e Lipase Kma,e (mmol) Vmaxb,e (lmol min 1 PC PLA1 HLL 1.36 ± 0.037 0.37 ± 0.0098 8841.73 ± 300.21 389.59 ± 9.56 5157.68 ± 128.54 227.26 ± 5.78 3780.51 622.29 TAG PLA1 HLL 1.53 ± 0.045 2.13 ± 0.078 3891.05 ± 87.98 8354.22 ± 261.32 2269.78 ± 56.89 4873.294 ± 134.87 1488.10 2283.10 mg 1 ) kcatc,e (s 1 ) kcat/Kmd,e (s 1 mM 1 ) Km: the substrate affinity constant. kcat: the turnover of the enzymatic reaction. Vmax: the maximal rate. kcat/Km: the catalytic efficiency. The values are mean ± standard deviation based on three independent experiments. Fig. 3. Variations in hydrolytic activity of (A) triolein, and (B) DOPC, with surface pressure by monolayer technique. PLA1 and TLL were injected into the reaction compartment of a zero-order trough (volume, 2.5 mL; surface area, 12.56 cm2). Buffer: 10 mM PBS, pH 7.0. Activities are expressed as the number of moles of substrate hydrolyzed per time (min) unit (M) and surface unit (cm2). pronounced lag phase of approx. 24 min was observed. This may indicate an energy barrier, due to conformational changes in the enzyme associated with the interfacial activation [8,51]. As expected, these results are in accordance with the kinetic parameters measured by the emulsion method. These findings support the idea that the C-terminal domain is involved in interactions with the PC interface. As PC is a major component of lipid membranes, we hypothesize that the C-terminal extension acts as a membrane anchor that increases the affinity of PLA1 towards phospholipid membranes. The amino acid sequence of C-terminal region present in PLA1 has high homology with those of GZEL and YLLIP8. In previous studies, it has been suggested that YLLIP8 is associated with Y. lipolytica cell wall as a membrane-bound protein [15]. The above evidence may render this hypothesis more attractive. Asp294 of the C-terminal helix and Arg87 of the a3 helix in PLA1 lid were found to be conserved among the RML, TLL and GZEL. The a9 helix of C-terminal region may control the opening of lid to regulate the activity of TAG by transiently interacting with the a3 helix of lid, following the similar ‘double-lock’ activation mechanism of GZEL [40]. We propose that the C-terminal region of PLA1 plays a significant role in interfacial activation to affect the chain-length specificity and the C-terminal helical tail anchoring onto PC to regulate the phospholipase activity. This hypothesis is in agreement with the observation that the C-terminal helical region of YLLIP8 acts as a transient membrane anchor [15]. Further experiments are required to validate this hypothesis, but it would be helpful to explain the role of an extra C-terminal region of PLA1 and bring new insight into rational design of novel lipases with intriguing functionalities. 4. Conclusion Acknowledgments The C-terminal region present in the lipase family is important for interfacial binding [51], signal transduction [52] and membrane-trafficking and -disruption [53]. In the present study, the extra C-terminal region of PLA1 was found to be essential for its substrate specificity and enzymatic property. The residue This work was supported by National High Technology Research and Development Program of China (2014AA093514 and 2014AA093601), National Science Funds Foundation of China (31471690), and Science and Technology Planning project of Guangdong province (2014B020204003 and 2015B020231006). 154 R. Xin et al. / Journal of Colloid and Interface Science 488 (2017) 149–154 Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jcis.2016.10.058. References [1] R. Aravindan, Lipase applications in food industry, Ind. J. Biotechnol. 6 (2007) 141–158. [2] F.I. Khan, B. Nizami, R. Anwer, K.-R. Gu, K. Bisetty, M.I. Hassan, et al., Structure prediction and functional analyses of a thermostable lipase obtained from Shewanella putrefaciens, J. Biomol. Struct. Dyn. 1102 (2016) 1–13. [3] A. Aloulou, J.A. Rodriguez, S. Fernandez, D. van Oosterhout, D. Puccinelli, F. Carrière, Exploring the specific features of interfacial enzymology based on lipase studies, Biochim. Biophys. Acta Mol. Cell Biol. Lipids 1761 (2006) 995– 1013. [4] A.M. Brzozowski, U. Derewenda, Z.S. Derewenda, A model for interfacial activation in lipases from the structure of a fungal lipase-inhibitor complex, Nature 351 (1991) 491–494. [5] M. Schweiger, G. Schoiswohl, A. Lass, F.P.W. Radner, G. Haemmerle, R. Malli, et al., The C-terminal region of human adipose triglyceride lipase affects enzyme activity and lipid droplet binding, J. Biol. Chem. 283 (2008) 17211– 17220. [6] I. Panaiotov, M. Ivanova, R. Verger, Interfacial and temporal organization of enzymatic lipolysis, Curr. Opin. Colloid Interface Sci. 2 (1997) 517–525. [7] R. Verger, F. Pattus, G. Pieroni, C. Riviere, et al., Regulation by the ‘‘interfacial quality” of some biological activities, J. De Chimie Physique Et De Physico Chimie Biologique 84 (1985) 673–681. [8] A. Karray, Z. Zarai, Y. Gargouri, R. Verger, S. Bezzine, Kinetic properties of pancreatic and intestinal sPLA2 from chicken and mammals using the monomolecular film technique, J. Colloid Interface Sci. 363 (2011) 620–625. [9] I. Jemel, A. Fendri, Y. Gargouri, S. Bezzine, Kinetic properties of dromedary pancreatic lipase: a comparative study on emulsified and monomolecular substrate, Colloids Surfaces B Biointerfaces 70 (2009) 238–242. [10] A. Fendri, A. Sayari, Y. Gargouri, Kinetic properties of Turkey pancreatic lipase: a comparative study with emulsified tributyrin and monomolecular dicaprin, Chirality 17 (2005) 57–62. [11] P. Calvez, S. Bussiéres, Éric Demers, C. Salesse, Parameters modulating the maximum insertion pressure of proteins and peptides in lipid monolayers, Biochimie 91 (2009) 718–733. [12] F.H. Wang, H. Zhang, Y.H. Liu, F.I. Khan, B. Yang, Y.H. Wang, Profiling substrate specificity of lecitase ultra to different kinds of phospholipids using monolayer technology, Eur. J. Lipid Sci. Technol. 118 (2016), http://dx.doi.org/10.1002/ ejlt.201600175 (accessed on 12 October 2016). [13] F. Carrière, C. Withers-Martinez, H. Van Tilbeurgh, A. Roussel, C. Cambillau, R. Verger, Structural basis for the substrate selectivity of pancreatic lipases and some related proteins, Biochim. Biophys. Acta - Rev. Biomembr. 1376 (1998) 417–432. [14] H. Chahinian, V. Belle, A. Fournel, F. Carrière, The role of pancreatic lipase C2-like domain in enzyme interaction with a lipid-water interface, Eur. J. Lipid Sci. Technol. 105 (2003) 590–600. [15] J. Kamoun, M. Schué, W. Messaoud, J. Baignol, V. Point, E. Mateos-Diaz, et al., Biochemical characterization of Yarrowia lipolytica LIP8, a secreted lipase with a cleavable C-terminal region, Biochim. Biophys. Acta - Mol. Cell Biol. Lipids 2015 (1851) 129–140. [16] J.G. Yang, Y.H. Wang, B. Yang, G. Mainda, Y. Guo, Degumming of vegetable oil by a new microbial lipase, Food Technol. Biotechnol. 44 (2006) 101–104. [17] G. Fernandez-Lorente, M. Filice, M. Terreni, J.M. Guisan, R. Fernandez-Lafuente, J.M. Palomo, LecitaseÒ ultra as regioselective biocatalyst in the hydrolysis of fully protected carbohydrates. Strong modulation by using different immobilization protocols, J. Mol. Catal. B Enzym. 51 (2008) 110–117. [18] Q. An, F. Wang, D. Lan, F.I. Khan, R. Durrani, B. Yang, et al., Improving phospholipase activity of PLA 1 by protein engineering and its effects on oil degumming, Eur. J. Lipid Sci. Technol. (2016) 1–9. [19] B. Yang, R. Zhou, J.G. Yang, Y.H. Wang, W.F. Wang, Insight into the enzymatic degumming process of soybean oil, JAOCS J. Am. Oil Chem. Soc. 85 (2008) 421– 425. [20] L.R. Gerits, B. Pareyt, H.G. Masure, J.A. Delcour, Native and enzymatically modified wheat (Triticum aestivum L.) endogenous lipids in bread making: a focus on gas cell stabilization mechanisms, Food Chem. 172 (2015) 613–621. [21] L.R. Gerits, B. Pareyt, J.A. Delcour, A lipase based approach for studying the role of wheat lipids in bread making, Food Chem. 156 (2014) 190–196. [22] R. Fernandez-Lafuente, Lipase from Thermomyces lanuginosus: uses and prospects as an industrial biocatalyst, J. Mol. Catal. B Enzym. 62 (2010) 197– 212. [23] G.P. McNeill, R.G. Ackman, S.R. Moore, Lipase-catalyzed enrichment of longchain polyunsaturated fatty acids, J. Am. Oil Chem. Soc. 73 (1996) 1403–1407. [24] R.C. Rodrigues, R. Fernandez-Lafuente, Lipase from Rhizomucor miehei as an industrial biocatalyst in chemical process, J. Mol. Catal. B Enzym. 64 (2010) 1–22. [25] M.M. Soumanou, U.T. Bornscheuer, Lipase-catalyzed alcoholysis of vegetable oils, Eur. J. Lipid Sci. Technol. 105 (2003) 656–660. [26] L.a. Nelson, T.a. Foglia, W.N. Marmer, Lipase-catalyzed production of biodiesel, J. Am. Oil Chem. Soc. 73 (1996) 1191–1195. [27] F. Hasan, A.A. Shah, A. Hameed, Industrial applications of microbial lipases, Enzyme Microb. Technol. 39 (2006) 235–251. [28] K. Bojsen, A. Svendsen, C. Fuglsang, S. Patear, K. Borch, J. Vind, A. Petri, S. Gladd, G. Budolfsen, G. Schroder, Novozymes A/S, Denmark. PCT International Application WO2000/32758, 2000. [29] D. Lan, Y. Tai, Y. Shen, F. Wang, B. Yang, Y. Wang, Efficient purification of native recombinant proteins using proteases immobilized on cellulose, J. Biosci. Bioeng. 113 (2012) 542–544. [30] X.L. Qin, H.H. Huang, D.M. Lan, Y.H. Wang, B. Yang, Typoselectivity of crude Geobacillus sp. T1 lipase fused with a cellulose-binding domain and its use in the synthesis of structured lipids, JAOCS J. Am. Oil Chem. Soc. 91 (2014) 55–62. [31] C. Gao, D. Lan, L. Liu, H. Zhang, B. Yang, Y. Wang, Site-directed mutagenesis studies of the aromatic residues at the active site of a lipase from Malassezia globosa, Biochimie 102 (2014) 29–36. [32] F. Corpet, Multiple sequence alignment with hierarchical clustering, Nucleic Acids Res. 16 (1988) 10881–10890. [33] P. Gouet, X. Robert, E. Courcelle, ESPript/ENDscript: extracting and rendering sequence and 3D information from atomic structures of proteins, Nucleic Acids Res. 31 (2003) 3320–3323. [34] W.L. DeLano, The PyMOL molecular graphics system, Schrödinger LLC Wwwpymolorg. Version 1 (2002). <http://www.pymol.org>. [35] K. Hill, C.B. Pénzes, B.G. Vértessy, Z. Szabadka, V. Grolmusz, É. Kiss, et al., Amphiphilic nature of new antitubercular drug candidates and their interaction with lipid monolayer, Prog. Colloid Polym. Sci. 135 (2008) 87–92. [36] B. Gzyl, M. Filek, A. Dudek, Influence of phytohormones on polar and hydrophobic parts of mixed phospholipid monolayers at water/air interface, J. Colloid Interface Sci. 269 (2004) 153–157. [37] L. Ding, J. Li, S. Dong, E. Wang, Supported phospholipid membranes: comparison among different deposition methods for a phospholipid monolayer, J. Electroanal. Chem. 416 (1996). [38] S. Ransac, I. Panaiotov, Monolayer techniques, Methods Mol. Biol. 286 (1997) 263–292. [39] U. Derewenda, L. Swenson, Y. Wei, R. Green, P.M. Kobos, R. Joerger, et al., Conformational lability of lipases observed in the absence of an oil-water interface: crystallographic studies of enzymes from the fungi Humicola lanuginosa and Rhizopus delemar, J. Lipid Res. 35 (1994) 524–534. [40] Z. Lou, M. Li, Y. Sun, Y. Liu, Z. Liu, W. Wu, et al., Crystal structure of a secreted lipase from Gibberella zeae reveals a novel ‘‘ double-lock ” mechanism, 1 (2010), pp. 760–770. [41] A.G. Marangoni, D. Rousseau, Engineering triacylglycerols: the role of interesterification, Trends Food Sci. Technol. 6 (1995) 329–335. [42] J.H. Lee, J.M. Son, C.C. Akoh, M.R. Kim, K.T. Lee, Optimized synthesis of 1,3-dioleoyl-2-palmitoylglycerol-rich triacylglycerol via interesterification catalyzed by a lipase from Thermomyces lanuginosus, N. Biotechnol. 27 (2010) 38–45. [43] N. Liu, Y. Wang, Q. Zhao, Q. Zhang, M. Zhao, Fast synthesis of 1,3-DAG by LecitaseÒ Ultra-catalyzed esterification in solvent-free system, Eur. J. Lipid Sci. Technol. 113 (2011) 973–979. [44] Y. Wang, M. Zhao, K. Song, L. Wang, S. Tang, W.W. Riley, Partial hydrolysis of soybean oil by phospholipase A1 (Lecitase Ultra), Food Chem. 121 (2010) 1066–1072. [45] G. Piéroni, Y. Gargouri, L. Sarda, R. Verger, Interactions of lipases with lipid monolayers. Facts and questions, Adv. Colloid Interface Sci. 32 (1990) 341– 378. [46] P. Reis, K. Holmberg, H. Watzke, M.E. Leser, R. Miller, Lipases at interfaces: a review, Adv. Colloid Interface Sci. 147–148 (2009) 237–250. [47] W. Tsuzuki, A. Ue, A. Nagao, M. Endo, M. Abe, Inhibitory effect of lysophosphatidylcholine on pancreatic lipase-mediated hydrolysis in lipid emulsion, Biochim. Biophys. Acta - Mol. Cell Biol. Lipids 1684 (2004) 1–7. [48] S. Laurent, M.G. Ivanova, D. Pioch, J. Graille, R. Verger, Interactions between b-cyclodextrin and insoluble glyceride monomolecular films at the argon/ water interface: application to lipase kinetics, Chem. Phys. Lipids 70 (1994) 35–42. [49] G. Pieroni, R. Verger, Hydrolysis of mixed monomolecular films of triglyceride/ lecithin by pancreatic lipase, J. Biol. Chem. 254 (1979) 10090–10094. [50] L. de La Fournière, M.G. Ivanova, J.P. Blond, F. Carrière, R. Verger, Surface behaviour of human pancreatic and gastric lipases, Colloids Surfaces B Biointerfaces 2 (1994) 585–593. [51] S. Bezzine, F. Carrière, J. De Caro, R. Verger, A. De Caro, Human pancreatic lipase: an exposed hydrophobic loop from the C-terminal domain may contribute to interfacial binding, Biochemistry 37 (1998) 11846–11855. [52] E.A. Nalefski, J.J. Falke, The C2 domain calcium-binding motif: structural and functional diversity, Protein Sci. 5 (1996) 2375–2390. [53] S.H. Gerber, J. Rizo, T.C. Südhof, The top loops of the C2 domains from synaptotagmin and phospholipase A2 control functional specificity, J. Biol. Chem. 276 (2001) 32288–32292.