Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease [1 ed.] 9781620813942, 9781620813669

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012. ProQuest

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

BIOCHEMISTRY RESEARCH TRENDS

LIPASE

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

FUNCTIONS, SYNTHESIS AND ROLE IN DISEASE

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Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

BIOCHEMISTRY RESEARCH TRENDS

LIPASE FUNCTIONS, SYNTHESIS AND ROLE IN DISEASE

HAMDI SASSI AND

SOFIEN CANNAMELA Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

EDITORS

Nova Science Publishers, Inc. New York

Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

Copyright © 2012 by Nova Science Publishers, Inc. All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers‘ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works.

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Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

CONTENTS Preface Chapter I

Chapter II

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Chapter III

vii Acquisition of New Properties under the Action of Lipases as a Biotechnological Tool Samia Soultani-Vigneron, Laurent Poisson, Françoise Ergan and Gaëlle Pencreac’h Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase in an Organic Solvent and their Functionalities Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi Solvent-Free Biocatalytic Synthesis of Polyglycerol Polyricinoleate (PGPR) Using Immobilised Candida Rugosa and Rhizopus Arrhizus Lipases S. Ortega, M. C. Montiel, M. F. Máximo and J. Bastida

Chapter IV

Fungal Lipases: Versatile Tools for Biocatalysis Selmene Ouertani and Habib Horchani

Chapter V

Secreted Phospholipase A2 Inhibitors from Cynara Cardunculus L. and Aloe Vera Extracts as Potential Therapeutic Drug for Inflammatory Diseases Sofiane Bezzine and Youssef Gargouri

Chapter VI

Chapter VII

Chapter VIII

Lipoprotein Lipase Activity in Sickle Cell Nephropathy: A Review Abiodun Mathias Emokpae Lipases in the Pharmaceutical Industry: An Approach for Racemic Drugs Fabiano Jares Contesini, José Valdo Madeira Junior, Gabriela Alves Macedo, Hélia Harumi Sato and Patrícia de Oliveira Carvalho Yeast Cell Surface Display of Lipases Evgeniya Y. Yuzbasheva, Tigran V. Yuzbashev, Ivan A. Laptev, Tatiana V. Vybornaya and Sergey P. Sineoky

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1

35

73 111

137

155

171

185

vi Chapter IX

Contents Microbial Lipases and their Applications in Structured Lipids José Valdo Madeira Junior, Fabiano Jares Contesini, Paula Speranza, Hélia Harumi Sato, Gabriela Alves Macedo and Patrícia de Oliveira Carvalho

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Index

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211

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PREFACE Lipases are ubiquitous enzymes which catalyze the hydrolysis of triacylglycerols to glycerols and free fatty acids. Due to their characteristics of enantioselectivity and regioselectivity and stability in organic solvents, they have gained importance in industrial applications. In this book, the authors examine the functions, synthesis and role in disease of lipases. Topics discussed include the action of lipases as a biotechnological tool; fungal lipases in biocatalysis; secreted phospholipase A2 inhibitors from cynara cardunculus L. and aloe vera extract as a potential therapeutic drug for inflammatory diseases; lipoprotein lipase activity in sickle cell nephropathy; lipases in the pharmaceutical industry; and yeast cell surface display of lipases. Chapter I - Lipases have been associated, these last three decades, with several benefits in enzymatic biotransformation. Indeed, the use of lipases to catalyze either the hydrolysis of various esters in aqueous media or the synthesis of esters in organic media make them essential in the implementation of enzymatic reactions in a wide range of applications such as in the food industry and cosmetics. This paper aims at establishing an overview of areas explored in order to convert byproducts into more added-value products and/or to improve the properties of common molecules by use of free and immobilized lipases. Among applications that take advantage of the hydrolytic ability of lipases, enzymatic hydrolysis of phospholipids from plant origin has been explored these last decades. The marine microalga Isochrysis galbana is one of these sources that have been used for docosahexaenoic acid enrichment via enzymatic hydrolysis. This part will particularly focus on the ability of lipases to selectively hydrolyze phospholipids from microalga in order to obtain a DHA-rich fraction. Application of lipases in synthetic reactions will be widely discussed and supported by studies in solvent-free-medium as well as organic solvents. In this case, lipase-catalyzed syntheses will be detailed and will concern: triglycerides, waxes, sugar esters and chlorogenic acid derivatives. This chapter will be an opportunity to shed light on the promising future of lipases. Chapter II - Lauroyl saccharides were synthesized at 50oC through the condensation in various organic solvents with various water contents using the immobilized lipase from Candida antarctica. The apparent equilibrium constants, KC, based on the concentrations of substrates and products could be correlated to the dynamic hydration numbers values of the saccharides, indicating that the water activity played an important role during the enzymatic

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viii

Hamdi Sassi and Sofien Cannamela

reaction in the microaqueous solvent. The KC values also depended on the kind of solvent, and was found to correlate well with the relative dielectric constant of the solvent. The condensation of octyl -D-glucoside and octanoic acid and the hydrolysis of 6-O-octanoyl glucoside using the immobilized C. antarctica lipase were carried out in acetonitrile under various conditions. The kinetics for these reactions could be expressed based on the pingpong bi-bi mechanism, and all the kinetic parameters were evaluated. The dependence of the activity on the water concentration was expressed by an empirical equation, which was useful for predicting the transient changes in the concentrations of the substrates and the products under any conditions. Acyl mannoses with chain lengths of 8 to 16 were continuously produced at 50oC using a plug flow reactor (PFR) packed with an immobilized lipase. The conversion of more than 50% was obtained at the superficial residence time equal to or longer than 20 min. The continuous production of acyl ascorbate was also carried out using a continuous stirred tank reactor (CSTR) or PFR at 50oC, and the productivity was ca. 6.0 × 10 for CSTR and 1.9 × 103 g/(L-reactor·d) for PFR for at least 11 days, respectively. The surface tensions of acyl saccharides, sugar alcohols and ascorbates in an aqueous solution were measured, and the critical micelle concentration, CMC, and the residual area per molecule were calculated. The CMC values were independent of temperature but dependent on the pH. Bacteriostatic activities of acyl sugar alcohols were examined. The number and orientation of the hydroxyl groups played important roles in the activity. Antioxidative properties of acyl ascorbates and alkyl ferulates against lipid in bulk and microcapsule systems were kinetically analyzed. The higher antioxidative activity was exhibited at the longer acyl or alkyl chain length of the ascorbates or ferulates. Chapter III - The use of lipases is continuously increasing due to its ability to catalyse esterification, interesterification, acidolysis, alcoholysis and aminolysis in addition to the hydrolytic activity on triglycerides, to produce industrially important products such as emulsifiers, surfactants, wax esters, chiral molecules, biopolymers, modified fats and oils, structured lipids, and flavour esters. The authors have developed the biocatalytic synthesis of a food additive named polyglycerol polyricinoleate (PGPR) and identified with the code E-476. PGPR is widely known as an excellent water-in-oil emulsifier in the food industry, because it forms very stable emulsions even when the water content is very high, such as 80%. Therefore, PGPR is used as emulsifier in tin-greasing emulsions for the baking trade, and for the production of low-fat spreads. However, the main application of PGPR is in the chocolate industry, where is used in the adjustment of rheological properties of chocolate, improving the moulding properties of the molten chocolate. An additional property of PGPR in chocolate is its ability to limit fat bloom. The enzymatic synthesis of PGPR by the catalytic action of one or more lipases (which act in mild reaction conditions of temperature and pressure, neutral pH and in a solvent-free system), makes the process environmentally friendly and avoids side reactions so that the obtained product has a higher purity and quality than the current marketed PGPR obtained by chemical processes. Chapter IV - Lipases are ubiquitous enzymes which catalyze the hydrolysis of triacylglycerols to glycerols and free fatty acids. Under certain experimental conditions, such as in the absence of water, they are capable of reversing the reaction. The reverse reaction leads to esterification and formation of Mono-, Di-, Tri-glycerides from fatty acids and glycerol. Due to their very interesting characteristics like high enantioselectivity and

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Preface

ix

regioselectivity and stability in organic solvents, lipases have gained in the last decade much more importance in industrial applications. Among these biocatalysts, fungal lipases represent an important family due to their low cost of extraction, thermal and pH stability, substrate specificity and activity in organic solvents. In fact, many fungal lipases have been produced, purified and biochemically characterized since the middle of the last century. The chief producers of commercial lipases are Aspergillus niger, Candida cylindracea, Humicola lanuginosa, Mucor miehei, Rhizopus arrhizus, Rhizopus delemar, Rhizopus japonicus, Rhizopus niveus and Rhizopus oryzae. These lipases are usually used in a variety of biotechnological fields such as food and dairy (cheese ripening, flavor development), detergent, pharmaceutical, agrochemical (insecticide, pesticide) and oleochemical (fat and oil hydrolysis, biosurfactant synthesis, polymer synthesis) industries… Chapter V - The aim of the present work is to evaluate the anti-inflammatory properties of Cynara cardunculus L. (Asteraceae) during its growth and Aloe vera leaf skin. Cynara cardunculus L. and Aloe vera leaf skin (AVLS) were extracted using various solvents. The anti-inflammatory activities of crude extracts were evaluated by measuring the inhibition potency of mammalian non pancreatic phospholipases A2 (hG-IIA). The methanol and acetone extracts of leaves of C. cardunculus L. harvested on February exhibit potent inhibition of hG-IIA (IC50 = 50 µg/ml and 70 µg/ml, respectively). However, the acetone extract of stems harvested on December inhibit the hG-IIA with a lower IC50 around 130 µg/ml. Fractionation on silica gel and hydrophobic gel of the methanol extract of leaves of C. cardunculus L. harvested on February increase the inhibitory effect and the IC50 riched 10 µg/ml. Meanwhile, the water extract of AVLS exhibits the highest inhibitory effect with an IC50 = 0.22 mg/ml and interestingly no effect was observed on the digestive phospholipase A2 (group IB) even at a concentration of 5 mg/ml. Antioxidant activities of AVLS were also analyzed and the most active extracts were observed when using chloroform ethanol (1/1) and ethyl acetate (IC50 = 0.274 and 0.326 mg/ml, respectively). Analysis of the total phenolic content reveals that the water extract of AVLS, with the best anti-PLA2 effect, was poor in phenolic molecules (2 mg GAE/g). A significant correlation was established between the total phenolic content of AVLS and the antioxidant capacity but not with the anti PLA2 activity. Chapter VI - The importance of lipoprotein lipase (LPL) in lipoprotein metabolism is well documented. It is a key enzyme in the regulation of fatty acid flux involved in the hydrolysis of triglyceride rich chylomicrons, very low density lipoprotein, intermediate density lipoprotein liberating free fatty acid and glycerol for uptake in the target tissue. The activity of this enzyme may be affected by race, environment, sex, genetic and disease conditions. Literature on LPL activities in sickle cell disease is scare. The objectives of this review are to highlight the variations in the activity of LPL in sickle cell disease, a chronic haemolytic disorder that manifests by a wide variety of clinical, biochemical and haematological features. Sickle cell disease(SCD) is the most common haemoglobin variant accounting for over 60% of the world‘s major haemoglobinopathies with 2-3 millions Nigerians affected. The biochemical features of sickle cell nephropathy (SCN) may be different from other nephropathies and the LPL activity in SCN, sex differences and it‘s activity in different haemoglobin genotypes are examined. Lipoprotein lipase activity in SCD patients appears to be different from sickle cell trait and normal haemoglobin in Nigerians. The biosynthesis of this enzyme may be down regulated in SCD due to subclinical inflammatory episodes and associated conditions. These

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Hamdi Sassi and Sofien Cannamela

altered enzyme activity and lipoproteins are reflected in changes in the composition and concentration of various lipoprotein fractions which may also lead to changes in lipid components of red blood cell membranes. Chapter VII - Microbial lipases are commonly applied in the hydrolysis of fats, with different industrial applications. However, under micro-aqueous conditions they catalyze synthetic reactions leading to the obtaining of enantiopure pharmaceuticals such as antihypertensive and anti-inflammatory drugs. Several chemically synthesized drugs such as ibuprofen, ketoprofen and atenolol are found as racemic mixtures. However, in most cases only one of the enantiomers presents the therapeutical properties, while the other isomer may be inactive or even toxic. Within this context, the use of lipases to enantiomerically catalyze the resolution of these pharmaceuticals or drug precursors has been shown as one of the most refined applications of this enzyme, due to the obtaining of such high added value products. Lipases with good stability, selectivity, mild operational conditions and broad substrate specificity can feasibly be produced by filamentous fungi, yeasts and bacterial strains, such as Aspergillus sp, Candida rugosa, Candida antarctica, Serratia sp and Pseudomonas sp. Of the papers found in the literature reporting this use of lipases, the lipase from an Aspergillus niger strain showed promising results in the enantiomeric resolution of racemic ibuprofen. This enzyme catalyzes the enantiomerical esterification of R-ibuprofen, resulting in a greater percentage of S-ibuprofen, the enantiomer that presents the therapeutic properties. The lipase from Candida antarctica was used in the kinetic resolution of racemic atenolol, the enantiomeric resolution of the atenolol being carried out by a transesterification reaction using vinyl acetate as the acylant agent and an organic solvent as the reaction medium. In addition, the lipase from Serratia marcescens was shown to enantioselectively hydrolyze chiral drug precursors, with interesting results. All of these features render the use of lipases attractive for biotechnological applications in the field of organic synthesis and in the pharmaceutical industry, with the expectation of a positive impact on the health of the population, which would be treated with more effective drugs. Chapter VIII - Yeast cell surface display has been studied extensively. Compared with the traditional immobilization methods, it does not include tedious purification processes, and therefore, it can be used for directed evolution of enzymes. A protein produced by a cell attaches to the cell surface immediately after being transported outside. In general, a gene encoding the target protein is fused with a nucleotide sequence encoding part or all of a cell wall protein. The obtained recombinant protein complex directs to the secretory pathway of the cell wall protein and incorporates with the cell wall by mechanism of anchorage of this cell wall protein. Typically, in yeast, the C-terminal part of a glycoprotein that contains a glycosylphosphatidylinositol (GPI) anchor attachment signal sequence is used for cell surface display. This system allows the protein to covalently bind to the β-1,6-glucans of the cell wall. A second method of protein display fuses the N-terminal part or the whole cell wall protein to the N-terminus of a target protein. Then, the protein attaches to the cell wall either by covalent linkage to β-1,3-glucans, by linkage through disulfide bridges to glycoproteins covalently attached to β-1,6-glucans, or by non-covalent adhesion. This system is preferable if the active site of the immobilized enzyme is located near the C-terminus. Such an enzyme is lipase. In this chapter, the authors briefly review the general approaches to yeast cell surface display and the current results of lipase-cell wall immobilization. The authors also describe the development of lipase display on cells of Yarrowia lipolytica. Y. lipolytica is an aerobic hemiascomycetous yeast of great interest, because of its capacity to produce high

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Preface

xi

levels of homologous and heterologous proteins. It naturally secretes several lipases, among them the mainly synthesized is Lip2. Six putative GPI-anchored cell wall proteins have been identified in silico, in addition to the already characterized glycoprotein Y. lipolytica Ylcwp1. Lip2 translation fusion with the C-termini of these proteins revealed that all proteins were capable of immobilizing lipase in the active form on the cell surface. One of these proteins presumably plays a role in the adhesion process. It was assumed that the C-domain of this protein covalently attaches to the cell wall, while its N-domain, containing repetitive amino acid motifs, may be responsible for non-covalent cell-cell incorporation by adhesion. Thus, an N-terminal system of cell surface display has been successfully developed. It was demonstrated the high level of cell-bound lipase activity which was comparable to the best results obtained with the secreted form of this enzyme. Moreover, the stability of the displayed Lip2 was significantly higher than that of the free enzyme. Therefore, the cell wallimmobilized Lip2 could be suitable for use as a whole cell catalyst, thereby providing a new low-cost substitute for free lipases. Chapter IX - Microbial lipases are industrially important enzymes since they present interesting biochemical properties, great selectivity for the substrate and catalyze different reactions such as hydrolysis, esterification, transesterification, epoxidation and ring-opening polymerization, amongst others. One important target for lipase application is fat modification, since the nutritional and biological functions of food lipids are mainly dependent on the chain length and degree of unsaturation of the fatty acids. Thus one possibility is to modify the lipids by incorporating a new fatty acid, or restructuring them by changing the positions of the fatty acids, resulting in structured lipids. Structured lipids may provide the most effective means of delivering the desired fatty acids for nutritional or health prevention purposes, such as the prevention of coronary diseases. These compounds are suitable for use as nutraceuticals, since their structure can be manipulated to suit specific patient requirements, and the specificity of lipases is probably the main attribute for the synthesis of structured lipids. An immobilized lipase from Rhizopus oryzae was reported to present high incorporation of oleic acid at sn-1,3 positions, in short reaction times. The lipase from Thermomyces lanuginosus (Lipozyme TLIM) catalyzed transesterification reactions to obtain vegetable oils rich in medium chain fatty acids and their hypolipidemic and hypocholesterolemic effects were evaluated. The lipase from Rhizomucor miehei was applied to obtain structured lipids for parenteral nutrition, and it is possible to use Brazilian sardine oil in the acydolysis of soybean fatty acids. The lipase from Rhizopus oryzae showed high incorporation of oleic acid at the extreme positions of triacylglycerols maintaining palmitic acid at the sn-2 position. This work obtained lipids identical to those of human milk by enzymatic acidolysis. The potential for commercialization of the enzymatic process for the large-scale synthesis of structured lipids exists, and needs to be explored to obtain benefits in nutritional, medical and food applications.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

In: Lipase Editors: Hamdi Sassi and Sofien Cannamela

ISBN 978-1-62081-366-9 © 2012 Nova Science Publishers, Inc.

Chapter I

ACQUISITION OF NEW PROPERTIES UNDER THE ACTION OF LIPASES AS A BIOTECHNOLOGICAL TOOL Samia Soultani-Vigneron, Laurent Poisson, Françoise Ergan and Gaëlle Pencreac’h Laboratoire « Mer, Molécules, Santé » EA Université du Maine, Institut Universitaire de Technologie de Laval, Calmette et Guérin, France

ABSTRACT Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

Lipases have been associated, these last three decades, with several benefits in enzymatic biotransformation. Indeed, the use of lipases to catalyze either the hydrolysis of various esters in aqueous media or the synthesis of esters in organic media make them essential in the implementation of enzymatic reactions in a wide range of applications such as in the food industry and cosmetics. This paper aims at establishing an overview of areas explored in order to convert byproducts into more added-value products and/or to improve the properties of common molecules by use of free and immobilized lipases. Among applications that take advantage of the hydrolytic ability of lipases, enzymatic hydrolysis of phospholipids from plant origin has been explored these last decades. The marine microalga Isochrysis galbana is one of these sources that have been used for docosahexaenoic acid enrichment via enzymatic hydrolysis. This part will particularly focus on the ability of lipases to selectively hydrolyze phospholipids from microalga in order to obtain a DHA-rich fraction. Application of lipases in synthetic reactions will be widely discussed and supported by studies in solvent-free-medium as well as organic solvents. In this case, lipasecatalyzed syntheses will be detailed and will concern: triglycerides, waxes, sugar esters and chlorogenic acid derivatives. This chapter will be an opportunity to shed light on the promising future of lipases. 

Corresponding author: [email protected]. Tel: +33-2-43-59-49-62, Fax: +33-2-43-59-49-58.

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1. INTRODUCTION Enzymes are natural catalysts found in all living organisms and are produced by biobased materials fermentation (Louwrier, 1998). Their peculiar properties, among which high chemo- and regioselectivities and ability to work under mild reaction conditions, are considered as major assets for their massive use in the synthesis and modification of natural products. Among industrial enzymes, the total output of lipases keeps increasing for the past few years as they are considered as a promising tool in biotransformation and biotechnology. Lipases (E.C.3.1.1.3), a subclass of esterases, catalyze the hydrolysis of ester linkages in long-chain triacylglycerols to release alcohol and acid moieties. The catalytic activity and stability of lipases depend on their three dimensional conformation and their hydration state. The lipase active site often involves a movable lid-region and lipase is active only at the interface between a hydrophobic solvent and an aqueous medium as this interaction is associated with the opening of the lid (Mala and Takeuchi, 2008; Gumel et al., 2011). Animals, plants and microorganisms are sources of lipases eventhough lipases from fungi and bacteria remain largely favored in industrial applications (Carriere et al., 1994; Bhardwaj et al., 2001; Olempska-Beer et al., 2006). Lipases specificities, organic solvent-tolerance and thermostability were exploited to multiply their industrial application in areas such as detergent industry, food industry, pulp and paper industry, oleochemical industry, bioconversion in aqueous media and organic synthesis (Sharma et al., 2001). Nowadays, the projected global lipase market by 2012 is estimated around 100 million USD. Current markets are dominated by a small number of manufacturers and most industrial applications concern detergent industry (41.2 %), sweeteners (21.4 %), dairy products (17.4 %), chemicals (7.3 %), pulp and paper (6.7 %), leather (6.3 %) and biofuels (0.1 %) (Figure 1). More specifically, the new way of using lipases in organic solvents in the 1980s (Zaks and Klibanov, 1984; Riva et al., 1988) provided an interesting approach and opened new horizons of experiencing a novel synthesis way to replace chemical reactions and to produce a whole range of molecules. Since then, lipases have found applications in reactions such as the regioselective acylation and deacylation of carbohydrates (Riva, 2001), acylation of digitonine (Danieli et al., 1999), esterification of ascorbic acid with a carotenoid (Humeau et al., 1998), ester synthesis from short and long-chain fatty acids (Coulon et al., 1995; Vulfson, 1994). This chapter does not aspire to cover exhaustively all of the research on the potential use of lipases but aims at presenting some selected examples based on our experience in this area. Particularly, lipase-catalyzed modification of natural products and lipase-catalyzed synthesis of new derivatives will be discussed. Lipase use in biofuel production, albeit it is considered as a promising biocatalyst in alternative energy strategies to convert vegetable oil into fuel, will not be discussed.

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Acquisition of New Properties under the Action of Lipases …

3

Source: BCC research 2007. Figure 1. Projected 2012 global lipase market.

2. LIPASE-CATALYZED MODIFICATION OF NATURAL PRODUCTS

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2.1. Lipase-Catalyzed Production of Lysophospholipids Lysophospholipids (LPLs) are glycerophospholipids (PLs) in which either the sn-1 position (1-LPL) or the sn-2 position (2-LPL) is not esterified with a fatty acid (Figure 2). In vivo, LPLs are the products of PL deacylation catalyzed by either phospholipases (A1 or A2) or lipases. LPLs play crucial biological roles, mainly as signaling molecules. They are also key intermediates in PLs remodeling since once produced they are reacylated by specific acyltransferases leading to novel PLs. Recently, some LPLs, particularly lysophosphatidic acid, have been shown to be biomarkers and potential therapeutic targets for human diseases. In addition, LPLs have many applications in food, cosmetic and pharmaceutical industries due to their emulsifying properties considered to be even higher than those of PLs. Numerous studies are dealing with the synthesis of LPLs via chemical and/or enzymatic approaches (D‘Arrigo and Servo, 2010). In this chapter, our interest will deal with the use of lipases as biocatalysts for the production of LPLs from natural PLs.

Figure 2. Structures of lysophospholipids. R: alkyl chain; X : polar group (hydrogen, choline, ethanolamine, serine or inositol).

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A. Natural Sources of Phospholipids Typically, natural PLs are isolated from animals or plants. In the case of PLs from plant origin, the major source of PLs is soybean lecithin (Patil et al., 2010). Lecithin is a mixture of various phospholipids in association with neutral lipids. It is usually recovered as a byproduct during soybean oil production. Crude lecithin is obtained by dehydration of the gum produced in the degumming process of the raw oil. Crude lecithin contains a minimum of 35 % oil, which is reduced to less than 2 % by repeated solvent extraction with acetone. Other readily available vegetable sources of PLs are sunflower (Penci et al., 2010) and canola (Pastuszewska et al., 2000). These three typical lecithins consist of three main phospholipid species: phosphatidylcholine (PC; 2040 %), phosphatidylethanolamine (PE; 20-30 %) and phosphotidylinositol (PI; 15-25 %). They contain mainly linoleic acid (40-60 %), oleic acid (10-25 %) and palmitic acid (1015 %) (Doig and Diks, 2003). In the case of PLs from animal origin, egg yolk represents the most readily available source of PLs. The PLs originated from egg yolk contain 80 % PC, 15 % PE and 1 % PI (Boselli and Caboni, 2000). As in PLs from plant origin, the three main fatty acids in PLs from egg yolk are oleic acid (30 %), palmitic acid (25-30 %) and linoleic acid (15-20 %). Arachidonic acid and docosahexaenoic acid (DHA), two healthy polyunsaturated fatty acids, amount to around 5 and 3 %, respectively (Gładkowski et al., 2011). Pure phospholipids, available from various suppliers, are purified mainly from soybean lecithin and egg yolk. Besides these well-known sources of PLs, some marine microalgae contain also PLs with composition of interest. Particularly, as shown in our laboratory, PLs from the Haptophyceae Isochrysis galbana Parke (strain CCAP 927/1) contain around 50 % of DHA. This ratio of DHA in I. galbana PLs is very high as compared to PLs from animals and plants as well as to PLs from other microalgae. Other fatty acids are myristic acid (25 %), plamitic and oleic acids (10 % each) (Devos et al., 2006). B. Lipase-Mediated Production of LPLs In addition to triacylglycerols, their natural substrates, lipases act on various other substrates, including PLs (Guo et al., 2005). They catalyze both the hydrolysis, acidolysis, alcoholysis and interesterification of the ester bonds of PLs (Vikbjerg et al., 2006). Moreover, they are readily available and do not need calcium as cofactor unlike phospholipases. In some studies, the production of LPLs from PLs uses a chemo-enzymatic approach. PLs are first subjected to an alkaline deacylation and LPLs are then produced by lipasecatalyzed esterification. For instance, Hong et al. (2011) described the lipase-catalyzed esterification of sn-glycero-3-phosphatidylcholine (GPC) originating from soybean PC and free conjugated linoleic acid. In this study, three commercially available immobilized lipases were compared to a phospholipase A1 and a phospholipase A2. In the reaction conditions used, the lipases were more effective than the phospholipases for the production of lysophosphatidylcholine (LPC). Among them, Novozym 435 (immobilized lipase from Candida antarctica, Novozymes, Dk) gave the highest yield (70 % under optimized conditions). Similarly, Virto and Adlercreutz (2000) performed the synthesis of LPC via transesterification of GPC and vinyl laurate catalyzed by Novozym 435. High conversions (>95 %) were readily achieved.

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Another way of synthesizing LPLs from PLs is to perform direct deacylation of either the sn-1 or the sn-2 position of original PLs using lipase-catalyzed reactions. Most lipases display regiospecificity towards the sn-1 position in PLs resulting in the production of 1-LPLs. Deacylation can be achieved by lipase-mediated alcoholysis leading to the simultaneous production of lysophospholipids and fatty acid esters as described by Ghosh and Bhattacharyya (1997). Yields around 70 % were achieved with the lipase from Mucor miehei. Deacylation can also be performed by lipase-mediated hydrolysis as, for instance, for the production of DHA-rich LPLs (Devos et al., 2006; Pencreac‘h et al., 2011). In this case, the initial PLs were DHA-rich PLs from the microalga I. galbana. For this purpose, a lipase able to release all the fatty acids from PLs except DHA was selected. Among 14 enzymes tested, Lipase F from Amano Enzyme Inc (Rhizopus oryzae lipase) fully met this objective (Figure 3). Indeed, the hydrolysis yields for myristic acid, palmitic acid and oleic acid were higher than 90 % whereas the hydrolysis yield for DHA was only 15 %. Lipase F gave good performances allowing production of LPLs containing 77 % of DHA.

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C. Characterization of Produced LPLs 1-LPLs are thermodynamically unstable as compared to 2-LPLs because of the presence of a primary alcohol group at the sn-1 position. Therefore, 1-LPLs can readily undergo spontaneous isomerization to 2-LPLs by non-enzymatic acyl migration especially under basic or alkaline conditions. The lipase-catalyzed hydrolysis of phospholipids leads mainly to 1LPLs as explained above. Consequently, under reaction conditions favoring acyl migration, 2LPLs issued from 1-LPLs are obtained. When solely 1-LPLs or 2-LPLs are required, acyl migration has to be carefully checked to obtain the desired product. To assess acyl migration during a deacylation process, different methods have been proposed but most of them used drastic chemicals or specific equipments such as LC-MS.

Figure 3. Hydrolysis of DHA-rich phospholipids by commercial lipase preparations. PLs from I. galbana were submitted to enzyme-catalyzed hydrolysis for 24h and released fatty acids were determined. Extent of release (hydrolysis yield) was expressed for each fatty acid relatively to the initial amount of the considered fatty acid in PLs before hydrolysis. Enzyme preparations were either from Amano Enzyme Inc, Japan (A: A. niger; AY: C. rugosa; F: R. oryzae; G: P. camembertii; M: M. javanicus; PS: B. cepacia; DF-15: R. oryzae R: P. roquefortii; Newlase F: R. niveus; Pancreatin F: Porcine pancreas) or from Novozymes, (Novozym 435: C. antarctica B; Lipozyme TL-IM: T. lanuginosa ; Lipozyme RM-IM: R. miehei). From Devos et al. 2006.

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In the case of DHA-rich LPLs production, an original multi-step enzymatic approach was used (Poisson et al., 2009). During the sn-1 deacylation using Lipase F, if no acyl migration from sn-2 to sn-1 occurs, the 1-LPLs produced remain unchanged. Then, a subsequent sn-2 specific deacylation using a phospholipase A2 (Lecitase 10 L from Novozymes, Dk) would release the fatty acids still esterified at the sn-2 position. However it was shown that Lecitase 10 L hardly releases free fatty acids suggesting that most acyl moieties actually migrated from the sn-2 to the sn-1 position and thus became inaccessible to Lecitase 10 L. Finally, in order to confirm the new sn-1 position of acyl moieties, Lipase PS (immobilized Pseudomonas cepacia lipase from Amano Enzyme Inc, Jp), which has been identified as non-regiospecific (Devos et al., 2006), was used to release free fatty acids from the LPLs. All fatty acids initially esterified onto PLs were actually recovered during this step, showing clearly that acyl migration does occur. Literature describes several methods to minimize acyl migration during deacylation such as the use of packed bed reactors (Mu et al., 2001) or the addition of borax buffer in the reaction medium (Morimoto et al., 1993). Reaction temperature may also be an important parameter to check (Yang et al., 2005). For instance, for the synthesis of DHA-rich LPLs, lowering the temperature from 40 °C to 30 °C during the reaction reduces DHA migration by 37 % (Poisson et al., 2009) However, even if acyl migration during the enzymatic reaction is restricted as much as possible, it may further happen during product purification or storage. Acyl migration can be completely prevented by chemically blocking the OH group at the sn-1 position. For instance, 1-acetyl-2-DHA-glycerophosphatidylcholine was synthesized by Lagarde and coworkers (2008). Acetylation was performed either by using acetic anhydride or by enzymatic transesterification.

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2.2. Lipase-Catalyzed Synthesis of Waxes Lipid upgrade from agricultural surplus or by-products as well as the recycling of lipid wastes such as waste cooking oil or activated sludge will be crucial for a sustainable development within the future decade (Hosseini and Ju, 2011; Huyng et al., 2011). In the same time, due to petrol depletion, a more and more important attention will be paid on energy consumption within industrial processes. In this area, enzymatic catalysis for the valorization of agro-resources nowadays constitutes the driving force of many research works.

A. What Is Wax? Waxes are a quite complex mixture of monoesters from long-chain fatty acids and longchain alcohols (fatty alcohol) with many others constituents which can vary greatly in proportions, such as triacylglycerols, primary and secondary alcohols, aldehydes and ketones. Because of their physico-chemical properties, waxes have been widely used for years as lubricants in many industrial products from shoe polishes to cosmetics. Waxes of commercial interest originate from a few natural sources. Before being prohibited, traditional source of wax esters was spermaceti, a white substance located in a cavity of the head of cetaceans. The deep-sea fish, orange roughy, is also a source of waxes but in depletion due to overfishing (Grigor et al., 1990).

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To construct their honeycomb, bees secrete wax that can be collected as a by-product during honey harvest. Nowadays, natural waxes are mainly extracted from the seeds of the shrub Simmondsia chinensis also called jojoba, which can be found in desert areas of southern California, Arizona and northern Mexico. Other vegetable waxes can also be harvested from the dry leaves of the Brazilian palm carnauba (Copernicia cerefira) (Christie, 2011)

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B. Ester Composition of Waxes To characterize waxes and to identify wax esters, different chromatographic methods can be used. Thin Layer Chromatography can be employed for a qualitative analysis with a classical solvent system of hexane/ethyl acetate/acetic acid (80:20:1 v/v/v) and visualization at 120 °C with a methanol sulphuric acid spraying (Huynh et al., 2011). Quantitative analyses of rather simple mixture of wax esters can be carried out by HPLC. Separation can be performed using a RP-18 column at 35 °C under isocratic conditions with a blend of acetone and acetonitrile as mobile phase (Poisson et al., 1999). For saturated wax esters, UV detection is not appropriate thus the monitoring must be carried out by using an Evaporative Light Scattering Detector or Mass Spectrometry. For a more detailed analysis, Gas Chromatography (GC) is the most widely used method. Due to the low volatility of certain wax esters, a rise of oven temperature to 350 °C is often necessary. In that case, CG columns stable at high temperature are required. Otherwise, wax esters should be hydrolyzed and derivatized prior to GC analysis. Using these separation techniques, ester composition of several natural waxes has been established. The major wax esters of jojoba are composed of monosaturated fatty acids and fatty alcohols i.e. C18:1, C20:1 and C22:1 as fatty acids, and C20:1, C22:1 and C24:1 as fatty alcohols. The main ester in orange roughy waxes is oleic acid oleyl ester (C18:1 as fatty acid and fatty alcohol) and palmitic acid cetyl ester (C16:0 as fatty acid and fatty alcohol) in spermaceti (Grigor et al., 1990; Christie, 2011). C. Wax Ester Production Since the end of the 80's, biotechnological works involving enzymatic engineering have attracted attention to produce wax esters of commercial interest (Mukherjee and Kiewitt, 1988). Figure 4 shows a general scheme of alcoholysis reaction where a lipase can catalyze wax ester synthesis from a triglyceride and a fatty alcohol as substrates. As erucic acid (C22:1n-9) was said to cause coronary disease, oils from rapeseed cultivars producing high amounts of this fatty acid (High Erucic Acid Rapeseed – HEAR) were no more allowed for human consumption and replaced by oils low in erucic acid like canola oil for instance. Research to develop the use of HEAR oil for non food applications has been engaged.

Figure 4. General scheme of alcoholysis reaction where substrates and products can be: A: triglyceride; B: fatty alcohol; C: wax ester; D: glycerol. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

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In this area, Trani et al. (1991) investigated the lipase-catalyzed production of jojoba wax substitute from HEAR oil and erucyl alcohol. The main feature of this work is the use of a solvent free reaction medium composed solely of substrates and enzyme allowing a reduced subsequent downstream processing. Indeed, using Lipozyme RM-IM (Novozymes, Dk) as catalyst and a reaction mixture composed of HEAR oil and a stoichiometric amount of erucyl alcohol (calculated from the fatty acid composition of HEAR oil), they manage to reach completion in 8 h and to produce the following esters resembling jojoba: linolenic acid erucyl ester, linoleic acid erucyl ester, oleic acid erucyl ester, 11-eicoasenoic acid erucyl ester and erucic acid erucyl ester. With a view to find possible applications to milk fat surplus, milk fat was used as raw material to produce waxes by lipase-catalyzed alcoholysis with oleyl alcohol (Poisson et al., 1999). Using the food grade Enzeco Lipase Concentrate preparation (EDC, USA) a reaction yield of 59 % was obtained with the production of four main wax esters: myristic acid oleyl ester, stearic acid oleyl ester, palmitic acid oleyl ester and oleic acid oleyl ester. Works also focused on heavy triglyceride fractions of sheep milk fat for the production of waxes with cetyl alcohol as co-substrate and Lipozyme RM-IM (Novozymes, Dk) as catalyst. Indeed, as sheep milk is rich in fat (two times richer than cow‘s milk) there is a great interest in its utilization for new purposes (Salis et al., 2003). To optimize the reaction yield, several parameters have to be carefully examined. First a screening of lipases is often necessary to determine the most convenient catalyst depending on the desired final product i.e. the chain length of the co-substrates and depending as well on the reaction implemented (alcoholysis, esterification...). Since physical properties of lipases and particularly their active conformation have been shown to vary with their hydration state, water control is certainly one of the most important factors influencing reaction rates even with alcoholysis, which does not involve water. In addition, since water greatly influences acyl migration, the amount of water can also influence conversion rates especially when using sn-1,3 regioselective lipases. Indeed, with this type of lipases, the evolution of conversion yields was shown to be inversely dependent on water activity (aw) whereas when working with a nonspecific lipase, an intermediate aw of 0.53 appears to be optimal (Salis et al., 2003). The co-product of wax synthesis i.e. glycerol, could accumulate around the enzyme and generate a barrier limiting the access of the substrates to the active site. To avoid the formation of this barrier the addition of silica gel to the reaction mixture have been proposed to trapped glycerol by adsorption as it is produced (Castillo et al., 1997). At last, temperature and substrate molar ratio constitute also two important parameters to monitor for an optimal wax ester production.

3. LIPASE-CATALYZED SYNTHESIS OF NEW DERIVATIVES 3.1. Lipase-Catalyzed Synthesis of Triglycerides in Solvent Free Media Tailor made triglycerides can be used to adjust the melting range of foods and cosmetics. In contrast to the chemically catalyzed esterification, the enzymatic ester synthesis already proceeds at moderate temperatures, thus avoiding the polymerization of unsaturated fatty

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acids. Another advantage is that the enzymatically-synthesized ester can usually be qualified as a natural ingredient. Triglycerides can be enzymatically synthesized from glycerol and fatty acids (Figure 5). When long-tail fatty acids are used, a two-phase system is present, a polar glycerol phase and a non-polar fatty acid phase. During esterification water is produced, the water produced accumulates in the polar phase, the esters accumulate in the non-polar phase. One way to enhance the production of triglycerides is the removal of the by-product water during synthesis. Lipase-catalyzed synthesis in organic solvents has been studied extensively for syntheses of esters, peptides and optically active compounds. Free or immobilized enzymes are used in two-phase, reversed phase and reversed micelles systems, where the organic substrates are dissolved in the solvent phase. However commercial scale-ups of such enzymatic syntheses have seldom been reported. One reason for that is that toxic and expensive solvents and/or surfactants are being employed that ought to be avoided, especially if the products are intended for use as foods or food ingredients. Lipase catalyses in solvent free systems have been reported in an attempt to make processes commercially feasible. The toxicity problem of solvents and surfactants is eliminated; the number of purification steps is reduced. However, applications are quite limited because of the high melting points of most organic substrates and the weak stability of enzymes at higher temperatures.

Figure 5. Multistep enzymatic-catalyzed synthesis of triglycerides. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

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A. Triolein Synthesis Immobilized Mucor miehei lipase catalyzes synthesis reactions between glycerol and oleic acid. No organic solvent is necessary to solubilize the substrates, which allows for the use of a reaction medium solely composed of the necessary substrates. Water produced in the reaction evaporates due to the high temperature used for the process. A conversion of 86 % of oleic acid into triolein is obtained when using the substrates in stoichiometric amounts. Varying the ratio of glycerol over oleic acid allows for the preferential synthesis of one of the glycerides. Some batch reactors have been set up using different means of removing the water: spontaneous evaporation, molecular sieves, vacuum, and dry air bubbling. This work was performed using Lipozyme RM-IM, a 1,3-regiospecific immobilized Rhizomucor miehei lipase catalyzing in theory synthesis of pure 1,3-diacylglycerol. The possibility of synthesizing triacylglycerol with a 1,3-specific lipase was explained by the phenomenon of spontaneous isomerization of the enzymatically formed 1-monoacylglycerol or 1,3-diacylglycerol into 2-monoacylglycerol or 1,2-diacylglycerol, respectively, followed by the enzymatic esterification of the free primary hydroxyl (Ergan et al., 1990). B. Synthesis of Medium Chain Triglycerides (MCT) Medium chain triglycerides (MCT) of caprylic acid (octanoic acid) and capric acid (decanoic acid) have been synthesized. MCT are used as nutritional supplement for patient suffering from malabsorption caused by intestinal resection or diseases and also as components of infant feeding formula. It is also used as a solvent or a carrier of lipophilic nutrients or drugs such as vitamin K and phospholipids, and as a base material for edible films or edible lubricants for foods and food processing. The commercial manufacturing process of such MCT is a direct esterification of medium-chain fatty acids and glycerol at high temperature and high pressure followed by alkali washing, steam refining, molecular distillation, ultrafiltration and activated carbon treatment for the purification of the product. Vacuum and dry air bubbling were applied for forced dehydration. When capric glyceride synthesis was conducted in a closed batch reactor, the final conversion was only 50 % at 40 °C while the open reactor gave a final conversion of 80 % under the same conditions. Adding a cold trap at the exhaust line of the reactor in an attempt to remove water more rapidly led to a 15 % increase in final conversion (Kim and Rhee, 1991). A yield (78 %) of tricaprylin synthesis can be achieved with a 1,3-specific immobilized lipase without any organic solvent. These experiments were conducted in relatively important volumes: the synthesis of more than 100 mL of glycerides was possible. Furthermore, the non-esterified acid and the traces of mono- and diglycerides can be easily eliminated by a simple passage of the warm (60 °C) reaction medium on a pure 70-230 mesh silica gel. The esterification conditions presented concern only batch conditions but such data may be very useful for the development of a continuous production of esters (Selmi et al., 1997a). Stoichiometric amounts of substrates are necessary and sufficient to achieve the best production of tricaprylin with free or silica gel adsorbed glycerol (Selmi et al., 1997b). C. Synthesis of N-3 Rich Triglycerides Fish oil is the most abundant and the cheapest source of Eicosapentaenoic acid (EPA, C20:5,ω3) and Docosahexaenoic acid (DHA, C22:6,ω3). Most fish oils do not contain more than a 30 % combined level of EPA and DHA. For example, the primary sources of most

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commercially used n-3 fish oil is anchovy (Engraulis ringens) and sardine (Sardinops sagax) oils that contain 15–22 % of EPA and 9–15 % of DHA. However, oils of concentration of 50– 90 %, with controlled ratios of EPA and DHA, are preferred for many supplement and pharmaceutical applications. Therefore, various groups have developed strategies for the manufacturing of highly concentrated EPA and DHA products. Traditional chemical esterification processes for triglyceride preparation from free fatty acids and monoester concentrates of the highly labile long-chain n-3 polyunsaturated acids are not really feasible either, as their all-cis n-3 framework will be partially destroyed by oxidation, cis-trans isomerization or double-bond migrations. They are also quite susceptible to polymerization as a result of the rather drastic conditions involved in terms of extreme pH and temperature. The use of lipases as catalysts offers a milder re-esterification method, resulting in less byproducts and better quality oils. Ocean Nutrition Canada Ltd. (Kralovec et al., 2012) has successfully converted ethyl esters (EE) to triglycerides (TG) both directly and via an intermediate hydrolysis step through the free fatty acid (FFA) form (EE-FFA first and then FFA-TG) using Novozym 435. Reesterification from FFA concentrate was significantly faster than that from EE and could be carried out at lower temperatures and gave products in higher yields. After routinely achieving 90 % conversion from FFA to TG with multiple re-use of the biocatalyst, the reaction was gradually scaled-up to manufacturing using proprietary packed enzyme bed reactors. A plant assembly of four reactors allows manufacturing up to 7500 kg of reesterified TG per day. The homogeneous triglyceride synthesis was accomplished by an immobilized nonregiospecific yeast lipase from Candida antarctica (Novozym 435 from Novozymes, Dk). Direct esterification of glycerol with stoichiometric amount of 99 % pure EPA or DHA as free fatty acids was conducted under vacuum (0.01- 0.1 Torr) at 65 °C in the absence of any solvent with 10 % dosage of lipase as based on the weight of substrates. The co-produced water was condensed into a liquid nitrogen cooled trap as the reaction proceeded, thus shifting the equilibrium toward completion. This reaction was observed to be superior to the corresponding interesterification of triacetin with n-3 enriched ethyl esters as well as the glycerolysis of n-3 enriched ethyl esters which were conducted under similar conditions. Furthermore, the Candida antarctica lipase was found superior to the immobilized 1,3regiospecific Mucor miehei lipase also provided by Novozymes (Dk) as Lipozyme RM-IM under these conditions. 97 % incorporation was obtained in 24 h, but after 72 h the reactions had proceeded to completion and reached 100 % conversion. The extended time required for the incorporation of the very last few percentages is noteworthy and warrants a comment. It is believed to be related to the purity of the substrates and the fact that an exact stoichiometric ratio was used in both reactions. This may be improved by using a slight excess (1 - 2 %) of the fatty acid substrates (Haraldsson et al., 1995). The synthesis of triglycerides by enzymatic esterification of polyunsaturated fatty acids (PUFA) with glycerol has been reported by other authors (Robles Medina et al., 1999). The lipase Novozym 435 from Candida antarctica was used to catalyze this reaction. The main factors influencing the degree of esterification and triglyceride yield were the amount of enzyme, water content, temperature and glycerol/fatty acid ratio. The optimum reaction conditions were established as: 100 mg of lipase; 9 ml hexane; 50 °C; glycerol/PUFA concentrate molar ratio 1.2/3; 0 % initial water; 1 g molecular sieves added at the start of reaction; and an agitation rate of 200 rpm. Under these conditions, a triglyceride yield of

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93.5 % was obtained from cod liver oil PUFA concentrate; the product contained 25.7 % eicosapentaenoic acid and 44.7 % docosahexaenoic acid. These optimized conditions were used to study esterification from a PUFA concentrate of the microalgae Phaeodactylum tricornutum and Porphyridium cruentum. With the first, a triglyceride yield of 96.5 %, without monoglycerides and very few diglycerides, was obtained after 72 h of reaction; the resulting triglycerides had 42.5 % eicosapentaenoic acid. A triglyceride yield of 89.3 % was obtained from a P. cruentum PUFA concentrate at 96 h of reaction, which contained 43.4 % arachidonic acid and 45.6 % EPA. These high triglyceride yields were also achieved when the esterification reaction was scaled up 5-fold.

D. Trierucin Synthesis Erucic (C22:1) acid has many potential or well-established uses, either in the form of glycerides or as free fatty acid. Erucic acid-rich oils have been employed as lubricants. They may also be used in the continuous casting of steel and in transmission oil fluids, or in the manufacture of cosmetic products through the synthesis of waxes that could be used as a jojoba oil substitute. The major erucic acid producing plants are rapeseed (Brassica napus), crambe (Crambe abyssinica), white mustard (Sinapis alba) and nasturtium (Tropaeolum). This latter species is the only one to contain trierucin. The lipase from Candida rugosa has been shown to discriminate against erucic acid. Advantage of this property has been taken to produce trierucin from high-erucic acid rapeseed (HEAR) oil. A method has been developed for extracting erucic acid from the oil as dierucin and subsequently enzymatically converting it to trierucin. Unrefined HEAR oil was hydrolyzed with lipase from C. rugosa to produce a mixture of free fatty acids and dierucin. Precipitation and filtration from cold ethanol gave 73 % pure dierucin, free of fatty acids. This dierucin was treated in two ways to produce trierucin. First, in the presence of an immobilized lipase and a known amount of water, some trierucin is produced by interesterification. Second, a more efficient route to trierucin utilized Rhizopus arrhizus lipase to completely hydrolyze dierucin to erucic acid, which was then combined with an appropriate amount of dierucin in the presence of an immobilized lipase to produce trierucin in a quantitative yield ((Trani et al., 1993). E. TG Synthesis: Effect of Chain Length and Unsaturation The triacylglycerol synthesis occurs more rapidly for the three longer fatty acids (tetradecanoic acid, hexadecanoic acid, and octadecanoic acid) than for decanoic acid and dodecanoic acid. In the first case after 6 h, almost 80 % of the initial fatty acids are esterified into triacylglycerol molecules while only 60 % of the dodecanoic acid and decanoic acid are converted to the related triacylglycerols. For all saturated fatty acids, the equilibrium is reached in less than 4 days. The higher yield of synthesized triacylglycerols is obtained for hexadecanoic acid and octadecanoic acids. At the end of the reaction, only triacylglycerols could be detected, leading to a calculated conversion yield of 100 %. In comparison, shorter chain fatty acids (C10–C14) present a lower conversion yield; the shorter the acyl chain is, the lower the yield becomes. Equivalent outcomes have been reported by the investigation of the influence of fatty acid chain length on triacylglycerol synthesis by Chromobacterium viscosum lipase in solvent-free medium. It can be suggested that medium chain fatty acids reaction media (C10–C14) are more polar than those with hexadecanoic acid or octadecanoic acid, leading to a lower rate of

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water elimination during the triacylglycerol synthesis. Furthermore, the different polarity of the different fatty acids could affect the rate of the acyl migration required for the reactions to go to completion. Another factor making a direct comparison of the different experiments difficult is the viscosity of the different reaction mixtures at 80 °C. It could be expected that the viscosity is very different, thereby affecting diffusion of fatty acid with mono- and diacylglycerol required for the reactions. The influence of unsaturation number for C18 on the reaction rates of triacylglycerols synthesis has also been studied. It is clearly shown that the number of unsaturations is responsible for a lower rate of triacylglycerol synthesis. Linoleic acid is esterified more slowly than oleic acid, which is itself esterified more slowly than octadecanoic acid. This is correlated with the rate of triacylglycerols synthesis. The possibility of synthesizing trilinolenin from linolenic acid and glycerol under these conditions has been tested. The esterification occurred; however, rapidly at this temperature (80 °C) problems of polymerization occurred, showing that it was not possible to compare the effect of zero and three double bonds in a solvent-free system. To compare the equilibrium position of saturated and unsaturated fatty acid, experiments were performed with saturated and unsaturated C18. In terms of yield at equilibrium, octadecanoic and oleic acids are completely converted to the related triacylglycerols while only 60 % of trilinolein is obtained when using linoleic acid. This indicates that a double bond in a fatty acid does not significantly affect the equilibrium. A two double-bond fatty acid presents a lower conversion yield. It can be suggested that the viscosity of the reaction mixture at 80 °C when using linoleic acid is very different, thereby affecting diffusion of fatty acid, mono- and diacylglycerol required for the reaction. A good yield of triacylglycerol synthesis by esterification of glycerol and C10–C18 fatty acids can be achieved with 1,3-specific immobilized lipase without any organic solvent. Such a simple system is proven to be useful for different acyl chain lengths. The best rates and equilibrium yields are obtained with long chain fatty acids. Indeed, a total esterification is obtained for hexadecanoic, octadecanoic, and oleic acids, leading to a complete synthesis of corresponding triacylglycerols with 100 % purity. In the reaction conditions presented here (i.e. 80 °C), decanoic acid is the shortest fatty acid that can be esterified with a good yield. Octanoic acid evaporates a little bit but also seems to have an important effect leading to enzyme inactivation. The particular interest of this work is the comparison of the effects of different types of fatty acids on the behavior of the enzyme (Selmi et al., 1998).

3.2. Lipase-Catalyzed Synthesis of Fatty Acid Sugar Esters (FASEs) Among the most studied enzymatic reactions in recent years, the lipase-catalyzed synthesis of FASEs in non-aqueous media has been widely discussed in the literature. These molecules are nonionic surfactants consisting of two groups: a water-soluble sugar as hydrophilic group and an acyl donor (fatty acids), soluble in oil and solvents, as lipophilic group. These esters are increasingly used as biosurfactants in food, pharmaceutical and cosmetic industries. Indeed, by varying the sugar nature and the fatty acid chain length, it is possible to obtain a set of molecules with a wide range of Hydrophilic-Lipophilic Balance (HLB) and therefore obtain interesting functional properties (foaming surfactants, fluidifier agents,

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Samia Soultani-Vigneron, Laurent Poisson, Françoise Ergan et al.

solubilizers, emulsifiers). They also show advantages in terms of biological activity (antibiotic), biocompatibility and biodegradability. Thus, they round off the range of nonionic surfactants commercially available and obtained by non-enzymatic chemical synthesis i.e. alkylglycosides and sucrose esters (Soultani et al., 2003). Despite a large number of papers dedicated to the enzymatic synthesis of FASEs, little industrial scale-up and production were achieved. This is mainly due, in the one hand, to difficulties to implement this kind of reactions in non-conventional media compared to chemical processes and on the other hand to the lack of knowledge of enzyme kinetics and mechanisms involved in heterogeneous and non-aqueous media. Indeed, most work to date has focused on the feasibility study without considering the constraints imposed by a possible industrialization of these enzymatic processes. Many lipases have been used to catalyze the synthesis of sugar esters. These enzymes are mostly from bacteria and fungi and are almost always used in an immobilized form with the best performances reported in literature for Novozym 435 and Lipozyme RM-IM. In industrial applications of FASEs, the immobilization of lipases has several advantages compared to the use of soluble lipases. Indeed, a better catalysis and stability of the lipase were observed. Thus, Redmann et al. (1997) have shown that, under the same operating conditions, the enzymatic synthesis of glucose fatty acid esters by direct esterification resulted in conversion yields of the acyl donor of about 55 % for immobilized lipases, whereas the reaction does not occur when using free lipases. The same result was obtained by Akoh and Mutua (1994) who explained the higher conversion yields by the high surface area available in the case of immobilized lipases compared to free enzymes. In addition, the immobilization of the enzyme usually leads to better thermal stability (Malcata et al., 1992). Finally, the immobilization carrier could allow the lipase to retain the amount of water needed for a proper active conformation (Garcia et al., 1996; Halling, 1994). The achievement of biocatalytic reactions in two-phase water-solvent media or organic solvent either pure or in mixture has been demonstrated (Laane et al., 1985; Zaks and Russell, 1988; Carrea et al., 1995; Sarney et al., 1997; Garcia-Alles and Gotor, 1998). The use of organic solvents can, however, have some disadvantages. As a matter of fact, with certain organic solvents, the enzyme can be denatured or inhibited causing a decrease of productivities and final yields. The first authors to be interested in the influence of organic solvents on the biocatalytic reactions were Brink and Tramper in 1985. These authors studied the influence of the polarity of the solvent on the enzyme activities using the Hildebrand solubility parameter (). This approach has been criticized by Laane et al. (1985) because of the lack of precision in the measurement and instead proposed to take into account the partition coefficient, log P as an indicator of the polarity of solvents. The same authors studied the effect of solvent hydrophobicity on the catalytic activity of biocatalysts from different sources. The hydrophobicity of the organic solvent thus has a significant effect on the catalytic activity of lipases with more or less important effects depending on the source of the enzyme. The study conducted by Bousquet et al. (1999) on the synthesis of unsaturated fatty acid glycoside esters catalyzed by Novozym 435 showed that the most suitable solvent for this type of synthesis is 2-methyl-2-butanol (2M2B) which log P value is 1.30. Indeed, this solvent yielded the highest fatty acid conversions (62 %) compared to other solvents. In most syntheses carried out with Candida antarctica lipase, it was found that the solvent used is the 2M2B (Faber et al., 1993; Coulon et al., 1999; Hugon, 2000). Indeed, this solvent is not a

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Acquisition of New Properties under the Action of Lipases …

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potential substrate for this lipase because it causes a steric hindrance at the active site (Ducret et al., 1995). According to Bousquet et al. (1999), it is a non-toxic solvent that can be used in areas such as pharmacology and cosmetology. In addition, 2M2B is widely used because it can dissolve the majority of substrates and does not cause lipase denaturation (Khaled et al., 1991). Regarding the acyl donor substrate, all lipases catalyze reactions with esters ranging from short (C4) to long chain fatty acids up to 22 carbons (Pleiss et al., 1998). However, it has been reported that each lipase shows a different specificity depending on the fatty acid. This specificity was explained by the shape of the active site, determined by molecular simulation, which varies from one enzyme to another. Concerning the second reaction substrate, although the chemical synthesis of FASEs is achieved especially with disaccharides, including sucrose, enzymatic synthesis of FASEs is usually achieved with monosaccharides. Indeed, the first enzymatic synthesis of FASEs in organic media using disaccharides has been carried out in 1988 by Riva et al. These authors achieved the synthesis of FASEs with three substrates: maltose, lactose and sucrose in dimethylformamide (DMF) at 45 °C with a protease from Bacillus subtilis as catalyst. In this case, the best conversion yield (96 %) was obtained with lactose. Thereafter, the most studied disaccharides were sucrose (Carrea et al. 1989; Dordick et al., 1993, Rich et al., 1995; Sarney et al., 1996) and to a lesser extent lactose (Sarney et al., 1994; Ku and Hang, 1995) and trehalose (Woudenberg van Oosterom et al., 1996; Hugon, 2000). One of the key challenges identified with enzymatic acylation of polysaccharides is their low solubility in hydrophobic solvents commonly used. However, many acylation reactions of monosaccharides or sugar alcohols have been carried out in recent years. The results reported in the literature showed that the nature of the sugar substrate plays an important role on the performance of enzyme-catalyzed synthesis of FASEs. Indeed, it determines the choice of the synthesis solvent and can also affect the properties of sugar esters as surfactants. Thus, it was reported that the more hydrophilic is the sugar, the lower is the air/water surface tension (Charlemagne and Legoy, 1995; Akoh, 1992). Similarly, Ducret et al. (1995) have shown that xylitol monooleate has a critical micelle concentration (CMC) 2 and 2.5 times lower than fructose and glucose monooleate respectively. These results suggest that hydrophobic interactions take place between sugar esters and water-soluble molecules of the medium. In conclusion, the choice of sugar is an important parameter not only in the performance of the reaction of enzymatic synthesis of FASEs but also in the final properties. By way of example let us take the case of the enzymatic synthesis of fatty acid fructose esters. This chapter shows that Novozym 435 catalyzed the synthesis of fructose esters with saturated fatty acids in a wide range of carbon chain lengths and concentrations at 60 ° C and 200 mbar in 2M2B. In this case, two reactions occur simultaneously with the production of water. Indeed, the first reaction results in monoesters, which in turn allow the synthesis of diesters in the second reaction (Figure 6). Many key parameters were demonstrated to affect the direct esterification synthesis of the fatty acid fructose mono- and diesters.

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A. Effect of the Initial Fatty Acid Concentration The consumption rate of the fatty acid varies between 1 and 6 mM.g-1.h-1 when the initial concentration of the fatty acid increases from 140 to 700 mM. However, it should be noted that when the stearic acid is used as the acyl donor, there is a drop in the consumption rate of the fatty acid when concentrations are above 560 mM. This profile has also been obtained by Erbeldinger et al. (1998) and Virto et al. (2000) during the Novozym 435 catalyzed synthesis of fatty acid sugar esters and fatty acid dihydroxyacetone esters. These results were explained on the one hand by a decrease in the substrates solubility in the reaction medium and on the other hand by diffusional limitations due to the increased medium viscosity. Increasing the fatty acid initial concentration between 140 and 700 mM causes a decrease in the fatty acid conversion yields from 80 to 20 % in the case of stearic acid after 50 hours of synthesis. In addition, when an excess of fructose is used compared to the fatty acid, the fatty acid conversion yield is improved but the reaction is slower. An increase in the initial fatty acid concentration improves also the conversion yield of fructose except for palmitic and stearic acids in concentrations above 560 mM for which there is a slight decrease of conversion yields. Furthermore, the same results were obtained by Mukesh et al. (1993) and Mutua and Akoh (1993) for Novozym 435 lipase-catalyzed synthesis of sugar esters. Indeed, this drop in the conversion yield was explained by the substrate inhibition of the lipase. This inhibition may be caused either by excessive acidification of the medium (Karra-Chaabouni et al., 1996) or by the formation of a hydrophobic barrier on the active site of the enzyme (Coulon et al., 1999). An increase of the initial fatty acid concentration causes a decrease in the selectivity toward monoesters. Similar results were reported by Ducret et al. (1995) and Hugon (2000) who showed that the equilibrium of the Novozym 435 catalyzed synthesis of sugar esters shifts in favor of diesters when an excess of the fatty acid is used. Lipase selectivity toward monoesters requires a large excess of fructose compared to the fatty acid. Indeed, 100 % monoesters were synthesized with 28 mM stearic acid and 5-fold fructose (140 mM).

Figure 6. Novozym 435-catalyzed synthesis of fructose mono- and diesters.

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Acquisition of New Properties under the Action of Lipases …

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An increase of the initial fatty acid concentration increases final total fructose esters. This result is explained in the literature by the mass action law. However, there was a drop in total fatty acid fructose esters with palmitic and stearic acids for concentrations above 560 mM. This result may be due to the enzyme inhibition by the substrate excess or by an accumulation of water produced during synthesis responsible of hydrolysis of the fructose esters formed.

B. Effect of the Initial Fructose Concentration The use of sugar substrate in the form of a solid phase has been reported to be an interesting approach for improving the reaction performance (Yan et al., 2001). In fact, suspended particles of the sugar continuously replenish the dissolved pool of the substrate as it is consumed by the reaction. However, substrates such as glucose have the potential to inhibit certain lipases, but not others (Tsuziki et al., 1999). The consumption rate of the fatty acid slightly increases when working with twice solid fructose. For instance, when stearic acid was used as the acyl donor, this rate varied from 3 to 3.5 mM.g-1.h-1 for 25 and 50 g.L-1 of fructose respectively. Therefore, it seems that when the fructose dissolution is a limiting factor, it leads to the decrease of soluble fructose in the medium, which causes a decrease in reaction rates. Similarly, an excess of sugar was used by Rich et al. (1995), Coulon et al. (1999) and Arcos et al. (1998) to improve the reaction rate. The fatty acid conversion yields were also improved when fructose concentrations increased. During the synthesis of fatty acid fructose esters, it was observed that using molecular sieves favors the synthesis of diesters. This effect may be offset by the gradual addition of fructose to the reaction medium. Indeed, as fructose is added, the reaction remains in favor of the monoesters. However, the initial fructose concentration had no significant effect on the final concentration of fatty acid fructose esters. Therefore, even when the dissolution of fructose is rate limiting, the final equilibrium is not affected. However, the addition of fructose in a fedbatch process has achieved concentrations of fatty acid fructose esters close to 120 g.L-1. C. Effect of the Fatty Acid Chain Length Consumption rate of the fatty acid did not differ significantly when the acyl donor chain length increased. Indeed, the variation does not exceed 1-2 mM consumed fatty acid.g-1.h-1 from caprylic acid (C8) to stearic acid (C18) depending on the initial fatty acid concentration. These results are in agreement with those obtained in the Novozym 435 lipase-catalyzed synthesis of esters of glucose and fatty acids from C4 to C20 (Degn et al., 1999) and those obtained during the synthesis of fatty acid trehalose esters from C5 to C18 (Hugon, 2000). However, these results are in contradiction with the structure representation of the active site of Candida antarctica lipase and its specificity reported by Pleiss et al. (1998). These authors highlighted the typoselectivity of this lipase towards C13 acyl donors. However, they also stated that the behavior of this lipase towards acyl donors longer than C13 could not be predicted. Indeed, in this case, the acyl donor can establish bonds with the hydrophobic surface of the lipase or withdraw within the catalytic pocket thereby altering the lipase specificity. The fatty acid and fructose conversion yields were similar regardless of the fatty acid except for stearic acid (C18) for which the highest conversion yields were obtained.

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Similar results were obtained by Cao et al. (1999) during the Novozym 435 catalyzed synthesis in acetone of glucose monoesters. Indeed, these authors have achieved the highest conversion yields with palmitic and stearic acids. This result was explained by the lipase preference for hydrophobic substrates and the precipitation of long carbon chain monoesters formed due to their low solubility in acetone. The proportion of monoesters, compared to total fructose esters, slightly decreases as the number of the acyl donor carbons increases. The same results were obtained in the synthesis of fatty acid trehalose esters (Hugon, 2000). For a given fatty acid concentration, increasing the length of the fatty acid carbon chain increases the production of fatty acid fructose esters. Thus, the concentration of the final fructose esters increased from 20 to 50 g.L-1 at 140 mM of caprylic acid (C8) and stearic acid (C18) respectively. This result can also be explained by the lipase activation in hydrophobic environments.

D. Effect of the Water Content The water content control of the medium, by using molecular sieves and by carrying out the synthesis under reduced pressure, improves the lipase activity and reactions are two times faster than when working under reduced pressure only. Similar results were obtained by Hugon (2000) who improved the reaction initial rates by decreasing the water content. In the presence of molecular sieves, the fatty acid conversion yields were improved. The same results were reported by Wehtje et al. (1997) who reported an increase of the conversion yield from 72 to 95 % before and after control of water by using molecular sieves. Similarly, Hugon (2000) has reported that the fatty acid conversion yields increase about 6-fold when the water content decreases from 0.1 to 0.01 %. Using molecular sieves during the enzymatic synthesis allows shifting the equilibrium toward sugar esters synthesis rather than hydrolysis. Thus, according to the equilibrium relationships, it is clear that the water control prevents the hydrolysis of monoesters that serve as substrates for the diesters synthesis. Therefore, the water removal from the medium is in favor of higher esters synthesis. Similarly, during the synthesis of dilauroyl 1,3 dihydroxyacetone, Virto et al. (2000) demonstrated that the water activity control enhanced diester concentrations. Hugon (2000) also reported that an increase in water content of the medium allowed shifting the equilibrium towards the synthesis of monoesters. Besides, reducing the water content in the medium by using molecular sieves is in favor of the production of fructose esters. Indeed, when water content is controlled, total fructose esters concentrations reach 60 to 70 g.L-1 depending on the fatty acid. This result can also be explained by the lipase activation in hydrophobic media. E. Effect of the Initial Hydrophobicity For a given solvent and at equal volumes, the initial hydrophobicity of the medium is provided primarily by the nature and concentration of the acyl donor. Thus, by varying the initial composition of the medium, the fatty acid consumption rate slightly changes and the more hydrophobic is the medium, the higher is the enzyme activity. However, when the hydrophobicity is above 1.3, there is a drop in these rates. This suggests that an initial hydrophobicity of 1.3 is necessary to achieve maximum enzymatic activity of the Novozym 435 lipase. However, one must consider the results reported by Humeau et al. (1998) who, for

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Acquisition of New Properties under the Action of Lipases …

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equal initial hydrophobicities, obtained different results depending on whether methyl palmitate is used alone or in mixture with hydrophobic solvents. In this case, the best results were obtained when methyl ester was used alone. Otto et al. (2000) have attributed this to the binding properties of the active site and the lipase-substrate interactions. The more hydrophobic is the medium, the less the reaction is selective toward the monoesters. These results are supported by those obtained by Virto et al. (2000) who favored the 1-monoacyl dihydroxyacetone synthesis in polar media and 1,3-diacyl dihydroxyacetone in non-polar media.

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3.3. Lipase-Catalyzed Synthesis of Polyphenol Derivatives Phenolic acids and flavonoids are the major dietary polyphenols. Phenolic acids include hydroxybenzoic acids, such as gallic acid, and hydroxycinnamic acids, such as caffeic acid, pcoumaric acid and ferulic acid. In plant material, the hydroxycinnamic acids are often in the form of glycoside or quinic esters such as chlorogenic acids. These are esters of caffeic, ferulic and quinic acids (Macheix et al. 2006) and have isomers as well as mono- or diacylated derivatives. The most common chlorogenic acid (CQA) is an ester of caffeic and quinic acids: 5-O-caffeoylquinic acid (5-CQA). Chlorogenic acids generate a growing interest both for their antioxidant and their pharmacological properties. Among the diacylated derivatives that are of medicinal value, the cynarin (1,3-O-dicafeoyl quinic acid) extracted from artichoke, is known for its hepatoprotective and cholagogue properties (Moglia et al., 2008). Similarly, 3-O-caffeoyl, 4dihydrocafeoyl quinic acid, isolated from a species of glasswort, Salicornia herbacea, shows an anti-tumor activity (Hwang et al., 2010). Chlorogenic acids also exist as glycosylated forms as 5-O-feruloyl quinic 4'-O-β-glycoside acid (Clifford et al., 2007), and more rarely as tri-acylated forms as 1,3,5-tri-O-(7,8-dihydrocafeoyl) quinic acid extracted from Podospermum laciniatum (Zidorn et al., 2005). This latter has a more important antioxidant activity compared to CQA due to the presence of three caffeoyl rings that increase this activity. Besides, tricaffeoyl quinic derivatives, in particular 3,4,5-tricafeoyl quinic acid, were demonstrated to have an anti-HIV activity (Tamura et al., 2006). Nowadays, the use of polyphenols is limited for sensitivity reasons and sometimes for their prooxidant properties, low bioavailability, their texture, astringency and unpleasant flavor. Moreover, a major difficulty in the industrial applications of polyphenols as antioxidants in lipid matrices is their hard incorporation due to their hydrophilic character. In order to increase the beneficial effects on health and resolve stability problems and incorporation of certain polyphenols in complex matrices, several methods have been proposed in the literature. This is the case of glycosylation (Dufour et al., 2002), lipophilisation (Figueroa-Espinoza and Villeneuve, 2005; Chebil et al., 2006; Villeneuve, 2007) or encapsulation (Fang and Bhandari, 2010). Polyphenol glycosylation can increase their antioxidant activity (Dufour et al., 2002). In contrast, lipophilisation change their hydrophilic character in order to increase their solubility in lipid media and improve their potential as antioxidants, both in food and in the body. Indeed, the esterification of the phenolic acids carboxyl group with fatty alcohols allows to increase the hydrophobicity and to synthesize multifunctional amphiphilic molecules (Figueroa-Espinoza and Villeneuve, 2005).

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Our interest will focus on the enzymatic lipophilisation of polyphenols. Indeed, biocatalysis allows considering the selective production of complex and original derivatives. Regarding lipophilisation of polyphenols, different types of enzymes were screened such as lipases, transferases, isomerases, esterases and proteases. However, lipases are the most frequently used for this type of biotransformation (Figueroa-Espinoza and Villeneuve, 2005; Chebil et al., 2006). The presence of a quinic moiety with non-phenolic hydroxyl groups and a carboxyl group makes the CQA one of the few polyphenols which allows to achieve the enzymatic synthesis of polyphenol derivatives either by alkylation or acylation (Figure 7). Knowing that much more data are reported in literature on enzymatic synthesis of polyphenol derivatives by alkylation, the data presented in this chapter examine only the reaction of acylation. The performance of polyphenol enzymatic acylation in terms of bioconversion yields and regioselectivity was demonstrated to depend on key-parameters as discussed below.

A. Effect of the Polyphenol Structure The nature of the polyphenol is a parameter that greatly influences bioconversion yields of the enzymatic acylation. For example, Kontogianni et al. (2001) have shown that when using palmitic acid methyl ester as the acyl donor, the rate of acylation of naringin is 10-fold higher than that of rutin. Under optimal conditions, i.e. with a acyl donor/naringin molar ratio of 7 and at 200 mbar, the bioconversion of naringin reached 92 %. The polyphenol structure was demonstrated to also affect the regioselectivity. Therefore, the highly regioselective acylation of naringin, described by Kontogianni et al. (2003), allowed the synthesis of a single product corresponding to the esterification of the only primary hydroxyl function of naringin. The study of the acylation reactions shows that Novozym 435 is regioselective of primary OH of glycosylated moiety of esculin as well as of the 4'''-OH rhamnose residue of rutin. The use of other lipases such as Pseudomonas cepacia or subtilisin esterases has also been reported but results in a lower regioselectivity (Chebil et al., 2006).

Figure 7. Enzymatic synthesis of polyphenol derivatives either by alkylation via the carboxyl group or by acylation via non-phenolic hydroxyl groups.

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Acquisition of New Properties under the Action of Lipases …

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Besides, Stevenson et al. (2006, 2007) undertook the direct acylation of phenolic compounds from crude extract of purified apple flavonoids using palmitic, cinnamic and phenylpropionic acids as acyl donors. The Novozym 435 used, showed a high regioselectivity for the acylation of primary OH sugars. However, no acylation occurred on the flavonoid aglycones or on the CQA. In all cases, the Novozym 435 catalyzed acylation of rutin and naringin with fatty acids is regioselective (Kontogianni et al., 2001, 2003; Ardhaoui et al., 2004; Mellou et al., 2005; Stevenson et al., 2006). When Novozym 435-catalyzed acylation occurs with a substrate having both primary and secondary hydroxyls, primary OH groups are preferred (Stevenson et al., 2007, Ardhaoui et al., 2004). Table 1 reports the transacylation regioselectivity of vinyl acetate depending on polyphenols (Lorentz, 2011).

B. Effect of the Solvent The nature of the organic solvents in terms of viscosity and polarity is a known parameter that influence lipase-catalyzed synthesis of polyphenol derivatives. The solvent must be inert toward the enzyme and must ensure good contact of the two substrates, which are of very different polarities. In order to express the effect in terms of solvent polarity, the most commonly used parameter is log P. Solvents with log P less than 3 as well as tertiary alcohols are good candidates. In addition, the use of organic co-solvents, ionic liquids or free-solvent medium is also described. Using a mixture of co-solvents has recently been described as an effective way to reconcile the solubility of the two substrates without using any chemical modification of the latter (Villeneuve, 2007). This method has been described for triglycerides acidolysis by phenolic derivatives. The influence of different solvents such as acetone, tetrahydrofuran (THF) or t-butanol was for example studied by Kontogianni et al. (2003). They achieved the naringin and rutin acylation with medium chain fatty acids (C8, C10, C12) in the presence of Novozym 435. When caprylic acid (C8) was used as the acyl donor, the highest conversion yields were observed in acetone, while for capric and lauric acids, t-butanol was demonstrated to be the best solvent. C. Effect of the Acyl Donor The acyl donor can influence the reaction depending on its nature (ester or acid), chain length and unsaturation. In the literature, the enzymatic transacylation of polyphenols was reported to be more efficient than the direct acylation in terms of bioconversion yields. Indeed, the direct acylation with fatty acids produces water that must be eliminated to reduce the hydrolysis reaction whether by using molecular sieves or by operating under reduced pressure while the transacylation with methyl, ethyl or vinyl esters was reported as irreversible As a matter of fact, Passicos et al. (2004) showed an increase in the bioconversion yields of naringin by palmitic acid (20 %) against methyl palmitate (90 %). However, in some cases, the Novozym 435-catalyzed transacylation gives poorer results than acylation. Indeed, Ardhaoui and coworkers (2004) showed that the transacylation is less efficient than the direct acylation because the hydrolysis of the acyl donor is predominant on the synthesis of flavonoid esters.

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Samia Soultani-Vigneron, Laurent Poisson, Françoise Ergan et al. Table 1. The transacylation regioselectivity of vinyl acetate depending on polyphenols (Lorentz, 2011)

Polyphenol Quercetin 7-OH

O

HO

OH

4'-OH

OH

3'-OH

4'-OH

OH O

2''-OH

OH

OH

HO O OH

Reference

6‘‘-O-acetate 3‘‘,6‘‘-O-diacetate For Novozym 435 6‘‘-O-acetate 3‘‘,6‘‘-O-diacetate 2‘‘,3‘‘,6‘‘-Otriacetate

Danieli et al. 1997 Chebil et al. 2007

Chebil et al. 2007

O

OH

Isoquercitrin

HO

Polyphenol derivative 4‘-O-acetate 3‘,4‘-O-diacetate 7,3‘,4‘triacetate

O

OH

3''-OH

OH

6''-OH

O

Rutin

For PS lipase 6‘‘-O-acetate 4‘,6‘‘-O-diacetate 4‘‘‘-O-acetate 3‘‘,4‘‘‘-O-diacetate

Danieli et al. 1997

6‘‘-O-acetate diesters

Katsoura et al. 2006

For QLG (Immobilized lipase from Alcaligenes sp.) 3-O-acetate 3,4‘diacetate Triacetate

Torres et al. 2010

OH O

HO

OH

OH

HO O OH

3''-OH

OH

O

O O

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H3 C HO

4'''-OH

HO OH

Naringin 6''-OH

OH

OH

O HO HO

O

O

O O

H3 C HO

OH

O

OH OH

Resveratrol 3-O H OH HO

OH

4'-O H

For Novozym 435 4‘-O-acetate 3,4‘diacetate

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Acquisition of New Properties under the Action of Lipases …

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Another approach using aromatic alcohols for the enzymatic synthesis of hydroxycinnamic esters has also been described. Torres de Pinedo et al. (2005) have reported the Novozym 435 catalyzed acylation in a series of orthophenolic di-alcohols with different fatty acids. The effect of the carbon chain length on the catalytic activity of lipases is widely described and is sometimes contradictory. Some studies have shown that Novozym 435 has a greater affinity towards short to medium chains resulting in increased catalytic performances (Pedersen et al., 2002). Similarly, Katsoura et al. (2006) reported that in the case of the rutin and naringin acylation with free fatty acids and their vinyl esters in ionic liquids, the highest bioconversion of 65 % was obtained for short chain lengths. Ardhaoui et al. (2004) also studied the effect of fatty acids from C6 to C18 on the rutin acylation by Novozym 435. They showed that the bioconversion yields increase until chain lengths of 12 carbons while no effect was observed for longer fatty acids. In addition, Salem et al. (2010) studied the influence of the carbon chain length of the acyl donor on the performance of the isoquercitrin acylation with Novozym 435. The influence of fatty acid chain lengths ranging from 4 to 18 carbons has been studied both in terms of conversion yields and initial reaction rates. It appeared that in all cases, thermodynamic equilibrium is reached between 48 and 72h and the conversion yield depends on the chain length. It varies from 66 % for ethyl butyrate to 38 % for ethyl stearate. Moreover, these authors showed that for chain lengths ranging from 4 to 12 carbons, the initial reactions rates were similar (~ 17x10-3 mmol.h-1) regardless of the fatty acid ester used. While for esters beyond 16 carbons, the initial reaction rates were maintained at around 10-2 mmol. h-1. Similarly, Mellou et al. (2005) reported that the acyl donor chain length did not affect the initial reaction rates for the naringin acylation neither with fatty acids nor their vinyl esters. Other authors, such as Bjorkling et al. (1989) showed a low Novozym 435 selectivity when the fatty acid chain length ranged from C8 to C18. Similarly, Kontogianni et al. (2001) have established no relationship between the acyl donor chain length and the conversion yields of rutin and naringin with Novozym 435 in various solvent systems (acetone, acetonitrile, THF). Otherwise, the study of the effect of unsaturated fatty acids leads to similar conclusions according to the authors. The effect of unsaturated fatty acids acylation of rutin catalyzed by Novozym 435 with C18 mono- and polyunsaturated fatty acids, was studied in acetone at 50° C by Mellou et al. (2005). It has been demonstrated that the unsaturation effect is not obvious, the bioconversions ranging from 70, 80 and 68 % for oleic acid (C18: 1), γ-linolenic acid (C18: 2), and α-linolenic acid (C18: 3) respectively. Similar results were reported by Viskupicova et al. (2010) for rutin acylation in 2M2B at 60 °C. The bioconversion yields reached 27, 29 and 29 % in 7 days with the same acyl donors against 34 % for stearic acid. In addition, Salem et al. (2010) also reported that the acyl donor unsaturation had no effect on the reaction kinetics of the isoquercitrin transacylation.

E. Effect of Temperature and Reduced Pressure Temperature is one of the most important key parameters in enzymatic synthesis. It affects both the viscosity of the medium, the enzyme activation or thermal denaturation as well as the substrates and products solubility (Chebil et al., 2006).

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Thus, increasing the reaction temperature reduced the time required to convert total 6''-Ophloridzin cinnamate by transesterification with Novozym 435 within 15 h at 60 °C, 4 h at 80 °C and 2 h at 100 °C (Enaud et al., 2004). These authors explained their results by the thermal activation of lipase and/or an easier removal of the co-product of the reaction, namely ethanol. Another explanation is a better distribution of phloridzin in the medium. In addition, these authors noted the perfect stability of the phloridzin and its product at high temperatures. Another example is the 1,1-phenyl-2-propanol transacylation with vinyl acetate and lipase from Pseudomonas cepacia, which led to bioconversion yields of 3 % at 60 °C and 39 % at 90 °C (Ema et al., 2003).

Figure 8. The reaction scheme of Novozym 435-catalyzed CQA acylation with palmitic acid in 2M2B.

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However, increasing temperature can affect the thermostability, resulting in the enzyme denaturation and a loss of activity. The literature states that this effect depends especially on the enzyme environment and the operating conditions. For example, Enaud et al. (2004) reported that 30 % of the Novozym 435 activity is lost after one day incubation at 80 °C in the 6''-O-phloridzin cinnamate synthesis. This was attributed both to the enzyme denaturation and possible interactions with the ethyl cinnamate. Similarly, Husson et al. (2008) reported that the catalyst, despite its immobilisation, lost 40 % of its activity after 8 h of reaction in the transacylation of 6-amino-1-hexanol in organic solvents. In contrast, Turner and Vulson (2000) showed that Novozym 435 catalyzed transesterification of octadecanol by methyl stearate without any loss of activity during 1 h at 130 °C. The study of Weitkamp et al. (2006), also showed that Novozym 435 is rather stable after 10 cycles of 4 h of use at 80 °C and 80 mbar for the synthesis of oleyl ferulate by transacylation of methyl ferulate. In addition, the acylation under reduced pressure may also improve the substrate-enzyme interactions and thus bioconversion yields. As an illustration, Figure 8 shows the reaction scheme of Novozym 435-catalyzed CQA acylation with palmitic acid in 2M2B achieved recently in our laboratory (Lorentz et al., 2010). Among the operating conditions tested for direct acylation synthesis of CQAmonoacylated derivatives of palmitic acid (C16), the best were obtained at 70 °C in the presence of a large excess of palmitic acid. These conditions resulted in a bioconversion yield of 95 % and a 4-O/3-O regioselectivity of 68 % in 10 days. It was demonstrated that the latter is significantly affected by the initial palmitic acid concentrations.

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CONCLUSION Lipases are becoming biocatalysts of choice for high-value biotechnological applications as they act on a variety of substrates and allow a large panel of reactions such as hydrolysis, interesterification, transesterification, acidolysis, aminolysis, etc. Nevertheless, the major drawback to the extensive use of lipases at an industrial scale is their low stability in their native state and their often prohibitive cost. Therefore, to overcome the lack of competitiveness of lipases compared to chemical catalysts and reduce the gap between the lab-scale and industry, academic research on lipases needs input of diverse disciplines and original tools and concepts have to be explored and deepened. Firstly, in order to cater to the needs of lipases in industries, novel lipase genes have to be isolated and the existing lipases are to be engineered for target properties. Thus, the modern methods of genetic engineering should allow extensive screening of new sources of microorganisms with a lipolytic activity and the development of molecular biology should enable researchers to tailor novel lipases with improved catalytic performances. As part of this objective, Godet et al. (2010) reported, for the first time, the isolation of a complete cDNA encoding a putative esterase from the marine microalga Isochrysis galbana. Secondly, many reaction mechanisms remain misunderstood and in a stage of hypothesis because of the lack of information regarding the active site of lipases and their specificity.

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The understanding of structure-function relationships between lipases and their substrates could be partially provided by molecular modeling. Indeed, computer-based modeling has recently been used to understand the mechanism and elucidate the regioselectivity of a few enzyme-catalyzed reactions. Finally, improvements of reaction yields and enzyme performances still need to be carried out. Although process engineering and reactor design modeling are increasingly used, advances are to be made to widespread the use of experimental designs in lipase-catalyzed reactions. Experimental design techniques should lead to the optimization of reaction conditions and a multivariable analysis allowing to comprehend the key-factors cross-effects on lipases activity. These tools should widen lipases applications in extreme conditions to best meet the industrial requirements and to take part in the emerging bioeconomy.

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REFERENCES Akoh, C. C. (1992) Emulsification properties of polyesters and sucrose ester blends I: Carbohydrate fatty acid polyesters. J. Am. Oil Chem. Soc. 69, 9-13. Akoh, C. C. and Mutua, L. N. (1994) Synthesis of alkyl glycoside fatty acid esters: Effect of reaction parameters and the incorporation of n-3 polyunsaturated fatty acids. Enzyme Microb. Technol. 16(2), 115-119. Arcos, J. A., Bernabé, M., Otero, C. (1998) Quantitative enzymatic production of 6-Oacylglucose esters. Biotechnol. Bioeng. 57, 505-509. Ardhaoui, M., Falcimaigne, A., Engasser, J. M., Moussou, P., Pauly, G., Ghoul, M. (2004) Acylation of natural flavonoids using lipase of Candida antarctica as biocatalyst. J. Mol. Catal. B: Enzym. 29(1-6), 63-67. Bhardwaj, K., Raju, A., Rajasekharan, R. (2001) Identification, purification and characterization of a thermally stable lipase from rice bran. A new member of the (phospho) lipase family. Plant Physiol. 127, 1728-1738. Björkling, F., Godtfredsen, E., Kirk, O. (1989) A highly selective enzyme-catalyzed esterification of simple glucosides. J. Chem. Soc., Chem. Commun. 934-935. Boselli, E. and Caboni, M. F. (2000) Supercritical carbon dioxide extraction of phospholipids from dried egg yolk without organic modifier. J. Supercrit. Fluids. 19, 45–50. Bousquet, M. P., Willemot, R. M., Monsan, P. and Boures, E. (1999) Enzymatic synthesis of unsaturated fatty acid glucoside esters for dermo-cosmetic applications. Biotechnol. Bioeng., 63(6), 730–736. Brink, L. E. S. and Tramper, J. (1985) Optimization of organic solvent in multiphase biocatalysis. Biotechnol. Bioeng. 27, 1258–1269. Cao, L., Bornscheuer, U. T., Schmid, R. D. (1999) Lipase-catalyzed solid phase synthesis of sugar esters. Influence of immobilization on productivity and stability of the enzyme. J. Mol. Catal. B: Enzym. 6(3), 279-285. Castillo, E., Dossat, V., Marty, A., Condoret, J. F., Combes, D. (1997) The role of silica gel in lipase-catalyzed esterification reactions of high-polar substrates. J. Am. Oil Chem. Soc. 74, 77-85.

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Carrea, G., Riva, S., Secundo, F. (1989) Enzymatic synthesis of various 1'-O-sucrose and 1O-fructose esters. Chem. Soc. Perkin Trans. 1, 1057-1061. Carrea, G., Ottolina, G., Riva, S. (1995) Role of solvents in the control of enzyme selectivity in organic media. Tibtech. 13, 63-70. Carriere, F., Thirstrup, K., Hjorth, S., Boel, E. (1994) Cloning of the classical guinea pig pancreatic lipase and comparison with the lipase related protein 2. FEBS Lett. 388, 6368. Chang, C.-S. and Wu, P. L. (2007) Synthesis of triglycerides of phenylalkanoic acids by lipase-catalyzed esterification in a solvent-free system. J. Biotechnol. 127(4), 694-702. Charlemagne, D. and Legoy, M. D. (1995) Enzymatic synthesis of polyglycerol-fatty acid esters in a solvent-free system. J. Am. Oil Chem. Soc. 72, 61-65. Chebil, L., Catherine, H., Falcimaigne, A., Engasser, J. M., Ghoul, M. (2006) Enzymatic acylation of flavonoids. Process Biochem. 41(11), 2237-2251. Chebil, L., Anthoni, J., Humeau, C., Gerardin, C., Engasser, J. M., Ghoul, M. (2007) Enzymatic acylation of flavonoids: Effect of the nature of the substrate, origin of lipase, and operating conditions on conversion yield and regioselectivity. J. Agri. Food Chem. 55(23), 9496-9502. Christie, W. W. (2011) Waxes structure, composition, occurrence and analysis. The lipid library. 2011, November 28, Available from: http://lipidlibrary.aocs.org/lipids/waxes/index.htm Clifford, M., Wu, W., Kirkpatrick, J., Kuhnert, N. (2007) Profiling the chlorogenic acids and other caffeic acid derivatives of herbal chrysanthemum by LC-MSn. J. Agric. Food Chem. 5(3), 929-936. Coulon, D., Girardin, M., Rovel, B., Ghoul, M. (1995) Comparision of direct esterification and transesterification of fructose by Candida antarctica lipase. Biotechnol. Lett. 17(2), 183-186. Coulon, D., Girardin, M., Ghoul, M., (1999) Enzymic synthesis of fructose monooleate in a reduced pressure pilot scale reactor using various acyl donors. Process Biochem. 34, 913-918. Danieli, B., Luisetti, M., Sampognaro, G., Carrea, G., Riva, S. (1997) Regioselective acylation of polyhydroxylated natural compounds catalyzed by Candida antarctica lipase B (Novozym 435) in organic solvents. J. Mol. Catal. B: Enzym. 3(1-4), 193-201. Danieli, B., Luisetti, M., Steurer, S., Michelitsch, A., Likussar, W., Riva, S., Reiner, J., Schubert-Zsilavecz, M. (1999) Application of lipase-catalyzed regioselective esterification in the preparation of digitonin derivatives. J. Nat. Prod. 62(5), 670-673. D'Arrigo, P. and Servo, S. (2010) Synthesis of lysophospholipids. Molecules. 15, 1354-1377. Degn, P, Pedersen, L. H., Duus, J.O, Zimmermann, W. (1999) Lipase-catalysed synthesis of glucose fatty acid esters in tert-butanol. Biotechnol. Lett. 21, 275-280. Devos, M., Poisson, L., Ergan, F., Pencreach, G. (2006) Enzymatic hydrolysis of phospholipids from Isochrysis galbana for docosahexaenoic acid enrichment Enzyme Microb. Technol. 39, 548-554. Doig, S. D. and Diks, R. M. (2003) Toolbox for exchanging constituent fatty acids in lecithins. Eur. J. Lipid Sci. Technol. 105, 359-367. Dordick, J. S., Hacking, A. J., Khan, R. A., (1993) Sucrose 6,4'-dicarboxylic esters. United States Patent n° 5 270 460.

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Samia Soultani-Vigneron, Laurent Poisson, Françoise Ergan et al.

Ducret, A., Giroux, A., Trani, M., Lortie, R. (1995) Enzymatic preparation of biosurfactants from sugar or sugar alcohols and fatty acids in organic media under reduced pressure. Biotechnol. Bioeng. 48, 214-221. Dufour, C., Da Silva, E., Potier, P., Queneau, Y., Dangles, O. (2002) Gallic esters of sucrose as efficient radical scavengers in lipid peroxidation. J. Agric. Food Chem. 50(12), 34253430. Ema, T., Kageyama, M., Korenaga, T. Sakai, T. (2003) Highly enantioselective lipasecatalyzed reactions at high temperatures up to 120 °C in organic solvent. Tetrahedron: Asymetry. 14, 3943–7. Enaud, E., Humeau, C., Piffaut, B., Girardin, M. (2004) Enzymatic synthesis of new aromatic esters of phloridzin. J. Mol. Catal. B : Enzym. 27,1–6. Erbeldinger, M., Ni, X., Halling, P. J. (1998) Enzymatic synthesis with mainly undissolved substrates at very high concentrations. Enzyme Microb. Technol. 23(1-2), 141-148. Ergan, F., Trani, M., André, G. (1990). Production of glycerides from glycerol and fatty acids by immobilized lipase in non-aqueous media. Biotechnol. Bioeng. 35,195-200. Faber, K., Ottolina, G., Riva, S. (1993) Selectivity-enhancement of hydrolase reactions, Biocatalysis. 8, 91–132. Fang, Z. and Bhandari, B. (2010) Encapsulation of polyphenols: a review. Trends Food Sci.Technol. 21(10), 510-523. Figueroa-Espinoza, M. C and Villeneuve, P. 2005. Phenolic acids enzymatic lipophilization. J. Agric. Food Chem. 53(8), 2779-2787. Garcia, T., Martinez, M., Aracil, G. (1996) Enzymatic synthesis of myristyl myristate. Estmation of parameters and optimization of process. Biocatal. Biotrans. 14, 67-85. García-Alles, L. F. and Gotor, V. (1998) Lipase-catalyzed transesterification in organic media: solvent effects on equilibrium and individual rate constants. Biotechnol. Bioeng. 59(6), 684-694. Ghosh, M. and Bhattacharyya, D. K. (1997) Enzymatic alcoholysis reaction of soy phospholipids. J. Am. Oil Chem. Soc. 74, 597-599. Gładkowski W., Kiełbowicz, G., Chojnacka, A., Gil, M., Trziszka, T., Dobrzanski, Z., Wawrzenczyk, C. (2011) Fatty acid composition of egg yolk phospholipid fractions following feed supplementation of Lohmann Brown hens with humic-fat preparations. Food Chem. 126, 1013–1018. Godet, S., Loiseau, C., Pencreac‘h, G., Ergan, F., Hérault, J. (2010) Isolation and sequence analysis of a cDNA encoding a novel putative esterase from the marine microalga Isochrysis galbana (Prymnesiophyceae, Haptophyta). J. Phycol. 46, 679-684. Grigor, M. R., Sutherland, W. H., Phleger, C. F. (1990) Wax-ester metabolism in the orange roughy Hoplostethus atlanticus (Beryciformes: Trachichthyidae). Marine Biology. 105, 223-227. Gumel, A. M., Annuar, M. S. M., Heidelberg, T., Chisti, Y. (2011) Lipase mediated synthesis of sugar fatty acid esters. Process Biochem. 46, 2079-2090. Guo, Z., Vikbjerg, A. F., Xu, X. (2005) Enzymatic modification of phospholipids for functionnal applications and human nutrition. Biotechnol. Adv. 23, 203-259. Halling, P. J. (1994) Predicting the behaviour of lipases in low-water media. Enzyme Microb. Technol., 16, 178–206

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Acquisition of New Properties under the Action of Lipases …

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Haraldsson, G. G., Gudmundsson, B. Ö., Almarsson, Ö. (1995) The synthesis of homogeneous triglycerides of eicosapentaenoic acid and docosahexaenoic acid by lipase. Tetrahedron. 51(3), 941-952. Hong, S. I., Kim, Y., Kim, C. T., Kim, I. H. (2011) Enzymatic synthesis of lysophosphatidylcholine containing CLA from sn-glycero-3-phosphatidylcholine (GPC)under vacuum. Food Chem. 129. 1-6. Hosseini, M. and Ju, L.-K. (2011) Use of algae to modify waste oil for biodiesel production, AOCS Meeting and Expo, 2011, December 2, Available from:http://www.aocs.org/ archives/am2011/abstracts.cfm?interest=Biotechnology Hugon, D. (2000) Production biotechnologique d‘une nouvelle gamme de tensioactifs: des esters de tréhalose et d‘acides gras. INPL thesis. Nancy (France). Humeau, C., Girardin, M., Rovel, B., Miclo, A. (1998) Effect of the thermodynamic water activity and the reaction medium hydrophobicity on the enzymatic synthesis of ascorbyl palmitate. J. Biotechnol. 63(1), 1-8. Husson, E., Humeau, C., Blanchard, F., Framboisier, X., Marc, I. Chevalot, I. (2008) Chemoselectivity of the N,O-enzymatic acylation in organic media and in ionic liquids. J. Mol. Catal. B: Enzym. 55(3-4), 110-117. Huynh, L. H., Do, Q. D., Kasim, N. S., Ju, Y. H. (2011) Isolation and analysis of wax esters from activated sludge. Bioresource Technol. 102, 9518-9523. Hwang, Y., Yun, H., Choi, J., Chun, H., Chung, Y., Kim, S., Kim, B., Kwon, K., Jeong, T., Lee, K., Jeong, H. (2010). 3-Caffeoyl, 4-dihydrocaffeoylquinic acid from Salicornia herbacea inhibits tumor cell invasion by regulating protein kinase C-δ-dependent matrix metalloproteinase-9 expression. Toxicol. Lett. 198(2), 200-209. Karra-Chaabouni, M., Pulvin, S., Touraud, D., Thomas, D. (1996) Enzymatic synthesis of geraniol esters in a solvent-free system by lipases. Biotechnol. Lett. 18, 1083-1088. Katsoura, M., Polydera, A., Tsironis, L., Tselepis, A., Stamatis, H. (2006) Use of ionic liquids as media for the biocatalytic preparation of flavonoid derivatives with antioxidant potency. J. Biotechnol. 123(4), 491-503. Khaled, N., Montet, D., Pina, M., Graille, J. (1991) Fructose oleate synthesis in a fixed catalyst bed reactor. Biotechnol. Lett. (13), 167-172. Kim, S. M. and Rhee, J. S. (1991) Production of medium-chain glycerides by immobilized lipase in a solvent-free system. J. Am. Oil Chem. Soc. 68(7), 499-503. Kontogianni, A., Sereti, V., Stamatis, H., Kolisis, F. (2001) Regioselective acylation of flavonoids catalyzed by lipase in low toxicity media. Eur. J. Lipid Sci. Technol. 103(10), 655-660. Kontogianni, A., Skouridou, V., Sereti, V., Stamatis, H., Kolisis, F. (2003) Lipase-catalyzed esterification of rutin and naringin with fatty acids of medium carbon chain. J. Mol. Catal. B: Enzym. 21(1-2), 59-62. Kralovec, J. A., Zhang, S., Zhang, W., Barrow, C. J. (2012) A review of the progress in enzymatic concentration and microencapsulation of omega-3 rich oil from fish and microbial sources. Food Chem. 131, 639-644. Ku, M. A. and Hang, Y. D. (1995) Enzymatic synthesis of esters in organic medium with lipase from Byssochlamys fulva. Biotechnol. Lett., 17, 1081-1084. Laane, C., Boeren, S., Vos, K. (1985) On optimizing organic solvents in muti-liquid-phase biocatalysis. Trends Biotechnol. 3, 251- 252.

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Samia Soultani-Vigneron, Laurent Poisson, Françoise Ergan et al.

Lagarde, M., Guichardant, M., Picq, M., Michaud, S., Doutheau, A. (2008) Method for preparing acetyl,docosahexaenoyl-glycerophosphocholine and use thereof for the delivery of polyunsaturated fatty acids. Patent WO/2008/068413. Lorentz, C. (2011) Nouveaux dérivés d‘acide cholorogénique: Synthèse enzymatique et valorisation. VENAM Thesis. Laval-Le Mans. France. Lorentz, C., Dulac, A., Pencreac‘h, G., Ergan, F., Richomme, P., Soultani-Vigneron, S.(2010) Lipase-catalyzed synthesis of two new antioxidants: 4-O- and 3-O-palmitoyl chlorogenic acids. Biotechnol. Lett. 3, 1955-1960. Louwrier, A. (1998) Industrial products – return to carbohydrate-based industries. Biotechnol. App. Biochem. 27, 1-8. Macheix, J. J., Fleuriet, A., Jay-Allemand, C. (2006) Composés phénoliques dans la plante: Structure, biosynthèse, répartition et rôles. In: Les polyphénols en agroalimentaire. TecandDoc Ed., Lavoisier, Paris. pp1-27. Mala, J. G. S. and Takeuchi, S. (2008) Understanding structural features of microbial lipasesAn overview. Anal. Chem. Ins. 3, 9–19. Malcata, F. X., Reyes, H. R., Garcia, H. S., Hill, C. G., Amundson, C. H. (1992) Kinetics and mechanisms of reactions catalysed by immobilized lipases. Enzyme Microb. technol. 14(6), 426-446. Mellou, F., Lazari, D., Skaltsa, H., Tselepis, A., Kolisis, F., Stamatis, H. (2005) Biocatalytic preparation of acylated derivatives of flavonoid glycosides enhances their antioxidant and antimicrobial activity. J. Biotechnol. 116(3), 295-304. Moglia, A., Lanteri, S., Comino, C., Acquadro, A., De Vos, R., Beekwilder, J. (2008) Stressinduced biosynthesis of dicaffeoyl quinic acids in globe artichoke. J. Agric. Food Chem. 56(18), 8641-8649. Morimoto, T., Murakami, N., Nagatsu A., Sakikibara, J. (1993) Regiospecific deacylation of glycerophospholipids by use of Mucor javanicus lipase. Tetrahedron Lett. 34, 24872490. Mu, H., Kurvinen, J-P., Kallio, H., Xu, X., Hoy, C. E. (2001) Quantitation of acyl migration during lipase-catalyzed acidolysis, and of the regioisomers of structured triacylglycerols formed. J. Am. Oil Chem. Soc. 78, 959-964. Mukesh, D., Jadhav, S., Sheth, D., Banerji, A. A., Newadkar, R. V., Behinakatti, H. S. (1993) Lipase catalysed reaction in a tubular reactor. Biotechnol. Tech. 7(8), 569-572. Mukherjee, K. D. and Kiewitt, I. (1988) Preparation of esters resembling natural waxes by lipase- catalyzed reactions. J. Agric. Food Chem. 36, 1336-1341. Mutua L. N. and Akoh C. C. (1993) Synthesis of alkyl glycoside fatty acid esters in nonaqueous media by Candida sp. lipase. J. Am. Oil Chem. Soc. 70, 43-46. Olempska-Beer, Z. S., Merker, R. I., Ditto, M. D., DiNovi, M. J. (2006) Food processing enzymes from recombinant microorganisms - A review. Reg. Toxicol. Pharmacol. 45, 144-158. Otto, R. T., Scheib, H., Bornscheuer, U. T., Pleiss, J., Syldatk, C., Schmid, R. D. (2000) Substrate specificity of lipase B: from Candida antarctica in the synthesis of arylaliphatic glycolipids. J. Mol. Catal. Part B. Enzym. 8, 201–211. Passicos, E., Santarelli, X., Coulon, D. (2004) Regioselective acylation of flavonoids catalyzed by immobilized Candida antarctica lipase under reduced pressure. Biotechnol. Lett. 26, 1073–1076,

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Pastuszewska, B., Jablecki, G., Swiech, E., Buraczewska, L., Ochtabinska, A. (2000) Nutritional value of rapeseed meal containing lecithin gums precipitated with citric acid. Anim. Feed Sci. Technol. 86, 117-123. Patil, V. V., Galge, R. V., Thorat, B. N. (2010) Extraction and purification of phosphatidylcholine from soyabean lecithin. Sep. Purif. Technol. 75, 138-144. Pedersen, N., Wimmer, R., Emmersen, J., Degn, P., Pedersen, L. (2002) Effect of fatty acid chain length on initial reaction rates and regioselectivity of lipase-catalyzed esterification of disaccharides. Carbohydrate Res. 337, 1179–1184. Penci, M. C., Constenla, D. T., Carelli, A. A. (2010) Free-fatty acid profile obtained by enzymatic solvent-free hydrolysis of sunflower and soybean lecithins. Food Chem. 120, 332–338. Pencreac'h, G., Ergan, F., Poisson, L. (2011) Production of lysophospholipids rich in DHA. Lipid Technol. 23, 250-252. Pleiss, J., Fischer, M., Schmid, R. D. (1998) Anatomy of lipase binding sites: the scissile fatty acid binding site. Chem. Phys. Lipids. 93, 67–80. Poisson, L., Jan, S., Vuillemard, J. C., Sarazin, C., Séguin, J. P., Barbotin, J. N., Ergan, F. (1999) Lipase-catalyzed synthesis of waxes from milk fat an oleyl alcohol. J. Am. Oil Chem. Soc. 76, 1017-1021. Poisson, L., Devos, M., Godet, S., Ergan, F., Pencreac‘h, G. (2009) Acyl migration during deacylation of phospholipids rich in docosahexaenoic acid (DHA): an enzymatic approach for evidence and study. Biotechnol. Lett. 31, 743-749. Redmann, I., Pina, M., Guyot, B., Blaise, P., Farines, M., Graille, J. (1997) Chemoenzymatic synthesis of glucose fatty esters. Carbohydrate Res. 300(2), 103-108. Rich, J. O., Bedell, B. A., Dordick, J. S. (1995) Controling enzyme-catalyzed regioselectivity in sugar ester synthesis. Biotechnol. Bioeng. 45, 426-434. Riva, S., Chopineau, J., Kieboom, A. P. G., Klibanov, A. M. (1988) Protease-catalyzed regioselective esterification of sugars and related compounds in anhydrous dimethylformamide. J. Am. Chem. Soc. 110, 584-589. Riva, S. (2001) Biocatalytic modification of natural products. Curr. Opin. Chem. Biol. 5, 106–111. Robles Medina, A., Esteban Cerdán, L., Giménez Giménez, A., Camacho Páez, B., Ibáñez González, M. J., Molina Grima, E. (1999) Lipase-catalyzed esterification of glycerol and polyunsaturated fatty acids from fish and microalgae oils. J. Biotechnol. 70(1-3), 379391. Salem, J., Humeau, C., Chevalot, I., Harscoat-Schiavo, C., Vanderesse, R., Blanchard, F., Fick, M. (2010) Effect of acyl donor chain length on isoquercitrin acylation and biological activities of corresponding esters. Process Biochem. 45(3), 382-389. Salis, A., Solinas, V., Monduzzi, M. (2003) Wax esters synthesis from heavy fraction of sheep milk fat and cetyl alcohol by immobilised lipases. J. Mol. Catal. B. 21, 167-174. Sarney, D. B, Kapeller, H., Fregapane, G., Vulfson, E. N. (1994) Chemo-enzymatic synthesis of disaccharide fatty acid esters. J. Am. Oil Chem. Soc. 71(7), 711-714. Sarney, D. B., Barnard, M. J., MacManus, D. A., Vulfson, E. N. (1996) Application of lipases to the regioselective synthesis of sucrose fatty acid esters. J. Am. Oil Chem. Soc. 73, 1481-1487.

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Sarney, D. B., Barnard, M. J., Virto, M., Vulfson, E. N. (1997) Enzymatic synthesis of sorbitan esters using a low-boiling-point azeotrope as a reaction solvent. Biotechnol. Bioeng. 54, 351–356. Selmi, B., Gontier, E., Ergan, F., Barbotin, J. N., Thomas, D. (1997a) Lipase-catalyzed synthesis of tricaprylin in a medium solely composed of substrates; water production and elimination. Enzyme Microb. Technol. 20(5), 322-325. Selmi, B., Gontier, E., Ergan, F., Thomas, D. (1997b) Enzymatic synthesis of tricaprylin in a solvent-free system: lipase regiospecificity as controlled by glycerol adsorption on silica gel. Biotechnol. Tech. 11(8), 543-547. Selmi, B., Gontier, E., Ergan, F., Thomas, D. (1998) Effects of fatty acid chain length and unsaturation number on triglyceride synthesis catalyzed by immobilized lipase in solvent-free medium. Enzyme Microb. Technol. 23(3-4), 182-186. Sharma, R., Chist, Y., Banerjee, U. C. (2001) Production, purification, characterization and applications of lipases. Biotechnol. Adv. 19, 627–662. Soultani, S., Ognier, S., Engasser, J. M., Ghoul, M. (2003) Comparative study of some surface active properties of fructose esters and commercial sucrose esters. Colloids Surf. A: Physicochem. Eng. Aspects. 227(1-3), 35-44. Stevenson, D. E., Wibisono, R., Jensen, D. J., Stanley, R. A., Cooney, J. M. (2006) Direct acylation of flavonoid glycosides with phenolic acids catalyzed by Candida antarctica lipase B (Novozym 435). Enzyme Microb. Technol. 39(6), 1236-1241. Stevenson, D. E., Parkar, S. G., Zhang, J., Stanley, R. A., Jensen, D. J., Cooney, J. M. (2007) Combinatorial enzymic synthesis for functional testing of phenolic acid esters catalysed by Candida antarctica lipase B (Novozym 435). Enzyme Microb. Technol. 40(5), 10781086. Tamura, H., Akioka, T., Ueno, K., Chujyo, T., Okazaki, K., King, P., Robinson, W. (2006) Anti-human immunodeficiency virus activity of 3,4,5-tricaffeoylquinic acid in cultured cells of lettuce leaves. Mol. Nut. Food Res. 50(4-5), 396-400. Torres de Pinedo, A., Penalver, P., Rondon, D., Morales, J. (2005) Efficient lipase-catalyzed synthesis of new lipid antioxidants based on a catechol structure. Tetrahedron 61(32), 7654-7660. Torres, P., Poveda, A., Jimenez-Barbero, J., Ballesteros, A., Plou, F. J. (2010) Regioselective lipase-catalyzed synthesis of 3-O-acyl-derivatives of resveratrol and study of their antioxidant properties. J. Agri. Food Chem. 58, 807-813. Trani, M., Ergan, F., André, G. (1991) Lipase-catalyzed production of wax esters. J. Am.Oil Chem. Soc., 68, 20-22. Trani, M., Lortie, R., Ergan, F. (1993) Enzymatic synthesis of trierucin from high-erucic acid rapeseed oil. J. Am. Oil Chem. Soc. 70(10), 961-964. Tsuzuki, W., Kitamura, Y., Suzuki, T., Mase, T. (1999) Effects of glucose on lipase activity Biosci. Biotechnol. Biochem. 63(8), 1467-1470. Turner, N. and Vulson, I. (2000) At what temperature can enzymes maintain their catalytic activity? Enzyme Microb. Technol. 27(1-2), 108-113. Vikbjerg, A. F., Rusing, J. Y., Jonsson, G., Mu, H., Xu, X. (2006) Strategies for lipasecatalyzed production and the purification of structured phospholipids. Eur. J. Lipid Sci. Technol. 108, 802-811. Villeneuve, P. (2007) Lipases in lipophilization reactions. Biotechnol. Adv. 25(6), 515-536.

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Virto, C. and Adlercreutz, P. (2000) Lysophosphatidylcholinesynthesis with Candida antarctica lipase B (Novozym 435). Enzyme Microb. Technol. 26, 630-635. Virto, C., Svensson, I., Adlercreutz, P. (2000) Candida antarctica lipase B-catalysed synthesis of dihydroxyacetone fatty acid esters. Biocatal. Biotransform. 18,13-29. Viskupicova, J., Danihelova, M., Ondrejovic, M., Liptaj, T., Sturdik, E. (2010) Lipophilic rutin derivatives for antioxidant protection of oil-based foods. Food Chem. 123(1), 4550. Vulfson, E. N. (1994) Industrial applications of lipases. In : Lipases – Their structure, biochemistry and application. Wooley, P. and Peterson, S. B. (Eds). Cambridge University Press, UK.pp271-288. Wehtje, E., Kaur, J., Adlercreutz, P., Chand, S., Mattiasson, B. (1997) Water activity control in enzymatic esterification processes. Enzyme Microb. Technol. 21, 502–510. Weitkamp, P., Vosmann, K., Weber, N. (2006) Highly efficient preparation of lipophilic hydroxycinnamates by solvent-free lipase-catalyzed transesterification. J. Agric. Food Chem. 54(19), 7062-7068. Woudenberg-van Oosterom, M., Van Rantwijk, F., Sheldon, R.A. (1996) Regioselective acylation of disaccharides in tert-butyl alcohol catalyzed by Candida antarctica lipase. Biotechnol. Bioeng. 49, 328–333. Yan, Y., Bornscheuer, U. T., Stadler, G., Lutz-Wahl, S., Reuss, M., Schmid, R. D. (2001) Production of sugar fatty acid esters by enzymatic esterification in a stirred-tank membrane reactor: Optimization of parameters by Response Surface Methodology. J. Am. Oil Chem. Soc. 78(2), 147-152. Yang ,T., Fruekilde, M. B., Xu, X. (2005) Suppression of acyl migration in enzymatic production of structured lipids through temperature programming. Food Chem. 92, 101107. Zaks, A., and Klibanov, A.M. (1984) Enzymatic catalysis in organic media at 100 C. Science. 224, 1249–1251. Zaks, A. and Russell, A. J. (1988) Enzymes in organic solvents: properties and applications. J. Biotechnol. 8(4), 259-269. Zidorn, C., Petersen, B., Udovicic, V., Larsen, T., Duus, J., Rollinger, J., Ongania, K., Ellmerer, E., Stuppner, H. (2005) Podospermic acid, 1,3,5-tri-O-(7,8 dihydrocaffeoyl) quinic acid from Podospermum laciniatum (Asteraceae). Tetrahedron Lett. 46(8), 12911294.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

In: Lipase Editors: Hamdi Sassi and Sofien Cannamela

ISBN 978-1-62081-366-9 © 2012 Nova Science Publishers, Inc.

Chapter II

SYNTHESIS OF ACYL SACCHARIDES AND ASCORBATES USING IMMOBILIZED LIPASE IN AN ORGANIC SOLVENT AND THEIR FUNCTIONALITIES Yoshiyuki Watanabe,*a, Takashi Kobayashib and Shuji Adachib a

Department of Biotechnology and Chemistry, Faculty of Engineering, Kinki University, Higashi-Hiroshima, Japan b Division of Food Science and Biotechnology, Graduate School of Agriculture, Kyoto University, Sakyo-ku, Kyoto, Japan

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ABSTRACT Lauroyl saccharides were synthesized at 50oC through the condensation in various organic solvents with various water contents using the immobilized lipase from Candida antarctica. The apparent equilibrium constants, KC, based on the concentrations of substrates and products could be correlated to the dynamic hydration numbers values of the saccharides, indicating that the water activity played an important role during the enzymatic reaction in the microaqueous solvent. The KC values also depended on the kind of solvent, and was found to correlate well with the relative dielectric constant of the solvent. The condensation of octyl -D-glucoside and octanoic acid and the hydrolysis of 6-O-octanoyl glucoside using the immobilized C. antarctica lipase were carried out in acetonitrile under various conditions. The kinetics for these reactions could be expressed based on the ping-pong bi-bi mechanism, and all the kinetic parameters were evaluated. The dependence of the activity on the water concentration was expressed by an empirical equation, which was useful for predicting the transient changes in the concentrations of the substrates and the products under any conditions. Acyl mannoses with chain lengths of 8 to 16 were continuously produced at 50oC using a plug flow reactor (PFR) packed with an immobilized lipase. The conversion of more than 50% was obtained at the superficial residence time equal to or longer than 20 min. The continuous production of

*Corresponding author: Department of Biotechnology and Chemistry, Faculty of Engineering, Kinki University, 1 Umenobe, Takaya, Higashi-Hiroshima 739-2116, Japan, E-mail address: [email protected]. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

36

Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi acyl ascorbate was also carried out using a continuous stirred tank reactor (CSTR) or PFR at 50oC, and the productivity was ca. 6.0 × 10 for CSTR and 1.9 × 103 g/(L-reactor·d) for PFR for at least 11 days, respectively. The surface tensions of acyl saccharides, sugar alcohols and ascorbates in an aqueous solution were measured, and the critical micelle concentration, CMC, and the residual area per molecule were calculated. The CMC values were independent of temperature but dependent on the pH. Bacteriostatic activities of acyl sugar alcohols were examined. The number and orientation of the hydroxyl groups played important roles in the activity. Antioxidative properties of acyl ascorbates and alkyl ferulates against lipid in bulk and microcapsule systems were kinetically analyzed. The higher antioxidative activity was exhibited at the longer acyl or alkyl chain length of the ascorbates or ferulates.

Keywords: Antioxidative surfactant; Continuous production; Equilibrium constant; Immobilized lipase; Reaction kinetics

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1. INTRODUCTION Saccharides and L-ascorbic acid are major hydrophilic components in food and essential for human life. Edible surfactant would be synthesized by chemical binding between the hydrophobic and hydrophilic components, and be usable as a food additive. Therefore, the condensation of saccharides or ascorbic acid with fatty acid has been exerted, and the improvement of efficiency for the reaction and various properties of the products have been studied [1-8]. Acyl saccharides synthesized in the reaction have been of much interest for use in many industries not only for food but also for chemicals, cosmetics and pharmaceuticals [9, 10]. Lipophilic derivatives of ascorbic acid acylated with a long-chain fatty acid has antitumor activity and metastasis-inhibitory effects [11, 12]. Since it has been recognized that medium-chain fatty acids or their glycerides enhance the absorption of hydrophilic substances in the intestine [13], it is expected that an ester of ascorbic acid and medium-chain fatty acid itself could be easily absorbed in the intestine with the physiological activity of vitamin C or that the ester could facilitate the absorption of hydrophilic substances. In addition, the oxidative stability of polyunsaturated fatty acid, which has important physiological functions such as antithrombotic, cholesterol depressant and antiallergenic properties [14, 15], may be improved by esterification of the acid with ascorbic acid. Acyl ascorbate seems to be promising emulsifiers with both surface-activity and reductivity. Alkyl ferulate and acyl kojic acid, which are synthesized through lipase-catalyzed condensation, would also have the similar properties, as ferulic and kojic acid is a compound with antioxidative activity. These products have been produced on an industrial scale based on chemical procedures. Lipases (EC 3.1.1.3) are widely used in the syntheses of various chemicals and polymeric materials [4]. Lipase-catalyzed reaction in aqueous system thermodynamically favors the hydrolysis, though the equilibrium of this reaction shifts towards condensation in nonaqueous organic solvent with low water content [16-22], in a solvent-free system [11, 23, 24] or under reduced pressure and/or in the presence of a desiccant [20, 25]. A certain amount of water is essential for hydration of lipase, even though water is not needed for condensation. The above-mentioned syntheses of acyl saccharide and ascorbate through lipase-catalyzed condensation would have some advantages; the simplicity of its reaction process, the moderate reaction conditions, and high regioselectivity of the enzyme. The reaction

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equilibrium constant is an important parameter to predict the equilibrium conversion for the synthetic reaction. The kinetics for the reaction is also required in order to elucidate the reaction properties. A continuous reaction would be preferred for their large-scale production to a batch system. Needless to say, the various functionalities, such as surface activity and antioxidative ability, of the products should be evaluated. In this chapter, the studies on the reaction equilibrium and kinetics for the condensation of saccharides or ascorbic acid with various fatty acids using immobilized lipase in an organic solvent, and the continuous production and functionalities of acyl saccharides and ascorbates were described.

2. FACTORS AFFECTING THE REACTION EQUILIBRIUM OF LIPASECATALYZED CONDENSATION

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2.1. Equilibrium Constants for the Lipase-Catalyzed Synthesis of Acyl Hexoses Lauroyl mannose was synthesized through the condensation in various organic solvents with various water contents using the immobilized lipase from Candida antarctica [26]. The reaction solvent was dehydrated over molecular sieves 5A in advance. A specified amount of water was added to the dehydrated solvent to adjust the initial water content of the solvent to a desired level, which was measured by Karl-Fischer titration. Mannose (0.25 mmol), lauric acid (1.25 mmol) and the immobilized lipase (100 mg) were placed in a vial. Five milliliters of the solvent was then added to dissolve or disperse the substrates. The vial was tightly screw-capped and then immersed in a thermo-regulated water bath at 50oC. Fifty microliters of the reaction mixture was removed and mixed with the same volume of 100 mmol/L 1octanol, which was used as the internal standard in the HPLC analysis, and twice the volume of the eluent. The concentration of the product was then determined. The equilibrium water content in the reaction mixture was also measured by Karl-Fischer titration. The product was identified to be 6-O-lauroyl mannose by 1H NMR. The produced lauroyl mannose was quantified using HPLC with an ODS column (4.6 mm×300 mm) and a refractometer. The eluent used was a mixture of acetonitrile and water (65/35 by vol.). The flow rate was 0.8 mL/min. The calibration curves were prepared using the products isolated from the reaction mixtures according to the reported methods [27] with a slight modification. For determination of mannose solubilized in various solvents, a Cosmosil 5NH2 column (4.6 mm×250 mm) was used. The eluent used was a mixture of acetonitrile and water (60/40 by vol.). The flow rate was 1.0 mL/min. The condensation was carried out in nine water-miscible organic solvents. Lauroyl mannose could be formed in acetonitrile, acetone, 2-methyl-2-propanol (tbutyl alcohol) and 2-methyl-2-butanol (t-amyl alcohol), but not in 2-propanol, propionitrile, dimethyl sulfoxide, N-methyl formamide and N,N-dimethyl formamide. Therefore, subsequent experiments were performed using the former four solvents. Di- and higher esters were not detected in the solvents [28]. Figure 1 shows the transient changes in the conversion of lauroyl mannose in the solvents with initial water contents of 0.029 to 0.043%. The conversion was defined as the molar ratio of the amount of lauroyl mannose to the initial amount of mannose. The conversion of every ester reached equilibrium within 3 days. The

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

Conversion [%]

60

40

20

0 0

20

40 60 Time [h]

80

100

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Figure 1. Transient changes in conversion of lauroyl mannose through immobilized-lipase catalyzed condensation at 50oC in (□) acetonitrile, (○) acetone, (▽) 2-methyl-2-propanol and (△) 2-methyl-2butanol with low water contents. The initial water contents of acetonitrile, acetone, 2-methyl-2propanol and 2-methyl-2-butanol were 0.039, 0.043, 0.029 and 0.029%(v/v), respectively.

conversion was the highest in acetonitrile and the lowest in 2-methyl-2-butanol, although the initial water content of each solvent was slightly different. The equilibrium conversions of lauroyl mannose in the above-mentioned four solvents with various initial water contents were higher at the lower initial water contents. This dependence of the conversion on the water content would be reasonable because water is one of the products in the condensation. However, the conversion largely depended on the kind of solvent. Among the solvents, acetonitrile gave the highest conversion at any initial water content, and a conversion of ca. 70% was realized in acetonitrile dehydrated with molecular sieves, the water content of which was 0.014% (v/v). In order to elucidate whether the reaction equilibrium constant, KC, itself was different among the solvents or other factors affected the yield, the KC values were calculated. The KC value was defined by Eq. (1) based on the equilibrium concentrations of the substrates and products.

KC 

CPeCWe CSeCFe

(1)

where C is the concentration in units of mol/L, and the subscripts S, F, P and W represent saccharide (mannose), fatty acid (lauric acid), product (lauroyl mannose) and water, respectively. The subscript e indicates equilibrium. The concentrations of product and water at equilibrium, CPe and CWe, were experimentally observed. The water content in units of %

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

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(v/v) was converted to the concentration in units of mol/L, assuming volume additivity among substrates, products and solvent. The molar volumes of the solvents were calculated from their densities and molecular masses. To obtain CSe, the solubility of mannose in each solvent with different water at 50oC was measured. The solubility depended on the kind of solvent. Mannose was the most soluble in 2-methyl-2-propanol and 2-methyl-2-butanol, followed by acetone and acetonitrile. In this reaction system, mannose was not fully dissolved in the solvent because of the limited solubility. Therefore, only the mannose dissolved in the solvent would be effective as a substrate for the condensation. When CS0－CPe, where CS0 is the overall initial concentration of mannose, was lower than the solubility CS at CWe, the concentration of mannose at equilibrium, CSe, was equal to CS0－CPe. On the other hand, when CS0－CPe > CS, the solubility CS at CWe was regarded as CSe. Lauric acid was completely dissolved in any case; therefore, its concentration at equilibrium CFe was calculated from CF0－CPe, where CF0 is the initial concentration of lauric acid. By substituting the concentrations of substrates and products at equilibrium into Eq. (1), the KC value was determined. The KC values were different by about two orders of magnitude among the solvents. The KC value was the highest in acetonitrile, intermediate in acetone, and small in 2methyl-2-propanol and in 2-methyl-2-butanol. The KC value evaluated here is an apparent one because it was defined based not on the activities but on the concentrations. The KC can be related to the intrinsic reaction equilibrium constant, Ka, which is defined based on the activities, by the following equation:

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KC 

γS γ F Ka γPγW

(2)

where  is the activity coefficient. If the water activity is only a parameter affecting the KC value, KC would be inversely proportional to W. The reaction system consisted of five components even if the immobilized enzyme was assumed to be inert, and it was too complicated to estimate the W value. Therefore, the W value under the assumption of a binary system consisting of water and organic solvent was estimated according to the UNIFAC [29]. However, the W did not correlate with the KC value. This might indicate that the water activity was not the sole factor affecting the reaction equilibrium in an organic solvent. Another possibility for the failure was that the W was not adequately evaluated because the assumption of a binary system was too simple. The dependence of the KC value on the equilibrium water concentration, CWe, was relatively weak for every solvent. Therefore, the KC values were averaged for each solvent to find a solvent parameter that could correlate with the averaged KC value and to obtain a criterion for selection of the solvent. There was a tendency for the KC to be smaller in a more hydrophobic solvent, but log P, where P is the partition coefficient between 1-octanol and water phases, did not seem to be a satisfactory parameter to correlate with the KC value. The Dimroth-Reichardt parameter for polarity of the solvents, ET(30), [30] did not correlate with the KC value at all. As shown in Fig. 2, the relative dielectric constant of the solvent, r, [31] correlated best with the KC (ln KC) value among the parameters tested. As a reason for the correlation, we guessed that the dissociation of lauric acid, which would be affected by the r value, was different among the solvents and that only the non-dissociated acid was effective for the condensation.

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40

Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi nDHN 14

15

16

17

18

19

5 4

lnKC

3 2 1 0 -1 -2

0

10

20 r

30

40

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Figure 2. Correlations of the apparent equilibrium constant, KC, for (□) lauroyl mannose synthesis in various solvents with the relative dielectric constants, r, of the solvents and for (○) lauroyl hexose in acetonitrile with the dynamic hydration number, nDHN, of hexoses.

The KC values for the syntheses of lauroyl glucose, galactose, mannose and fructose in acetonitrile at 50oC and various equilibrium water concentrations were also evaluated [28]. The KC value largely depended on the kind of hexose. Except for lauroyl fructose, there was a tendency that the KC was slightly higher at the lower CWe. The kind of solvent also significantly affected the KC value. When a saccharide hydrates, the water activity decreases. Therefore, it would be supposed that a hexose with stronger binding of water gives a larger KC value although the activities of other components would also be affected by the presence of the hexose. The dynamic hydration number, nDHN, of hexose was selected as a measure of the extent of hydration, and the relationship between the KC values observed in acetonitrile and the nDHN of hexoses [32] used as substrates was examined (Fig. 2). As the KC value depended on the CWe, the KC value averaged over all the CWe values for each product is plotted in the figure. As expected, there was a positive correlation between ln KC and nDHN. This indicates that the water activity plays an important role for the condensation in microaqueous organic solvents.

2.2. Effect of the Solvent on the Reaction Equilibrium in Tertiary Alcohol and Nitrile Various kinds of organic solvents are used for the synthesis of acyl saccharides by lipase. Some of them are tertiary alcohols, nitriles, and ketones. As described above, the equilibrium yield of the acyl saccharide was different in a different organic solvent, and the equilibrium constant based on the substrate concentration was also different. The equilibrium constant in this case could be quantitatively related with dielectric constant of an organic solvent.

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

41

Equilibrium constant

102

101

100 0

0.5

1.0

Volumetric fraction of nitrile

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Figure 3. Dependence of the equilibrium constant for the synthesis of 1-butyl decanoate in the mixture of tertiary alcohol and nitrile on the volumetric fraction of a nitrile. The open and closed symbols represent the mixture of 2-methyl-2-butanol and 2-methyl-2-propanol, respectively, with acetonitrile ( ), propionitrile ( ), and butyronitrile ( ).

In order to discuss the reaction equilibrium for the synthesis of esters by the lipasecatalyzed condensation more generally, the equilibrium constant for the formation of 1-butyl decanoate from 1-butanol and decanoic acid was estimated as a model reaction system [33]. Nitriles (acetonitrile, propionitrile, and butyronitrile), tertiary alcohols (2-methyl-2-propanol and 2-methyl-2-butanol), and their mixtures were used as a reaction medium for the estimation of equilibrium constant for the formation of 1-butyl decanoate. The equilibrium constant in nitriles was greater than that in tertiary alcohols. And it was 1.9 and 1.4 in 2methyl-2-propanol and 2-methyl-2-butanol, respectively, but it ranged from 21 to 38 in nitriles; i.e. the equilibrium constant did not depend greatly on the alkyl chain length of an organic solvent but depended greatly on the kind of its polar group (cyano or hydroxyl group). In addition, the equilibrium constant rose with increasing the volumetric fraction of a nitrile in a solvent mixture of nitrile and tertiary alcohol. The relationship between the equilibrium constant and the volumetric fraction of nitriles can be expressed by a curve (Fig. 3). These results suggests that the reaction equilibrium can be controlled to a certain degree by changing the composition of the reaction medium that was consisted of tertiary alcohols and nitriles. To relate a change of these equilibrium constants to a solvent parameter, the infrared (IR) spectra of decanoic acid and 1-butyl decanoate were measured in nitriles, tertiary alcohols and their mixtures. Strong absorption peak of the C=O double bond derived from an ester or a fatty acid was observed in the IR spectra. The absorption peak from decanoic acid shifted from 1713cm-1 to 1736cm-1 with increasing the volumetric fraction of a tertiary alcohol. Meanwhile, the change of the peak from 1-butyl decanoate was within 1720–1732cm-1

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

regardless of the fraction of a tertiary alcohol, and the variation was smaller than a case of decanoic acid. From these, it is suggested that the interaction between decanoic acid and tertiary alcohol was stronger than that between 1-butyl decanoate and tertiary alcohol. Such a difference in strength of the interaction causes the difference of the equilibrium constant in a different organic solvent. The wavenumber of the absorption peak of the C=O double bond of the decanoic acid could be roughly related to the equilibrium constant by an exponential function (Fig. 4), and the equilibrium constant abruptly increased with increasing the wavenumber. From these results, a property of the polar group of an organic solvent was connected quantitatively with the equilibrium constant.

2.3. Addition of Desiccant Because the condensation is thermodynamically controlled, removal of one or more of the products is effective for increasing the conversion. In the lipase-catalyzed condensation in an organic solvent, a desiccant such as molecular sieve is often added to the reaction system to remove water, one of the products. However, the criterion for the amount of the desiccant to be added to achieve the desired conversion had not been elucidated. In this context, we proposed a method for predicting the apparent equilibrium conversion in the presence of a molecular sieve based on the apparent reaction equilibrium constant, the adsorption isotherm of water onto the molecular sieve, the solubility of a hydrophilic substrate in the solvent, and the mass balance in terms of water (Fig. 5) [34]. The validity of the method was demonstrated for the lipase-catalyzed synthesis of monoacyl hexose in a water-miscible organic solvent [34].

Equilibrium constant

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102

101

100 1710

1720

1730

1740

Wavenumber of a C=O peak [cm-1] Figure 4. Dependence of the equilibrium constant for the synthesis of 1-butyl decanoate on the wavenumber of a C=O peak in an IR spectrum.

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

Reaction equilibrium constant based on concentrations KC 

C Pe C We CSe C Fe

Adsorption isotherm of water onto molecular sieve q We 

bC We 1  aC We

43

Solubility of sugar in reaction medium (empirical) S   exp( C We )

Mass balance of water VC W0  VC Pe  VC We  wqWe

Equilibrium conversion to ester (product) x Pe  C Pe / CS0

Figure 5. A method for predicting the equilibrium conversion for the synthesis of monoacyl hexose in the presence of molecular sieve.

The apparent reaction equilibrium constant, KC, is defined by Eq. (1). The solubility of hexose in the solvent, S, could be expressed empirically as a function of the concentration of water, CW, by Eq. (3).

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S   exp( CW )

(3)

where α and β are the constants, depending on the kinds of hexose and solvent, and the temperature. As shown in the inset of Fig. 6, the adsorption isotherm of water onto the molecular sieve was expressed by the Langmuir equation: qWe 

bC We 1  aC We

(4)

where qWe is the amount of water adsorbed, and a and b are the constants. The mass balance equation with respect to water is given at equilibrium by Eq. (5).

VC W0  VCPe  VC We  wqWe

(5)

where V is the volume of the solvent, w is the amount of the molecular sieve, and CW0 is the initial water concentration. The equilibrium product concentration, CPe, in the presence of the molecular sieve can be estimated by solving Eqs. (1), (4), and (5) simultaneously, and the equilibrium conversion, xe, is given by dividing CPe by CS0. Figure 6 shows the equilibrium conversion for the lauroyl mannose synthesis in 2-methyl-2-propanol and 2-methyl-2-butanol in the presence of various amounts of molecular sieve 3A. The solid curves were calculated by the proposed method and expressed well the experimental results.

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

0.4

qWe × 103 [mol/g]

Equilibrium conversion

0.8

10

5

0 0

0.05 0.10 CWe [mol/L]

0.15

0

0 50 100 Concentration of molecular sieve 3A [g/L]

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Figure 6. Equilibrium conversion at 50°C for the lauroyl mannose in () 2-methyl-2-propanol and ()2-methyl-2-butanol at different amount of molecular sieve 3A [34]. Inset: Adsorption isotherms of water onto molecular sieve 3A in 2-methyl-2-propanol and 2-methyl-2-butanol at 50°C.

Undesirable effects were also observed when molecular sieve 4A was added into the reaction system for the synthesis of oleoyl L-ascorbate in acetone [35]. The addition of molecular sieve 4A into the system increased both the initial reaction rate and the conversion. However, the conversion was gradually decreased on prolonged reaction time. The addition of excess amount of the molecular sieve lowered the conversion. These phenomena were ascribed to the adsorption of the product, oleoyl L-ascorbate, onto the molecular sieve and degradation of the product.

2.4. Improvement of the Product Concentration by the Use of a Sugar Derivative It is desirable to use a substrate at high concentration or to remove water to raise the product concentration in the lipase-catalyzed condensation from the viewpoint of the reaction equilibrium. However, it is very difficult to adopt the former one for the synthesis of acyl saccharides because of the poor solubility of a sugar in an organic solvent. The product concentration in the synthesis of acyl saccharide is generally low since the substrate concentration is low. Therefore some techniques to give the higher concentration of the sugar which is one of the substrates are proposed. For example, they are as follows: (1) addition of a sugar-solubilizing solvent such as dimethyl sulfoxide (DMSO) and pyridine to an organic solvent as an adjuvant, (2) using an ionic liquid as a reaction medium, and (3) raising the solubility of the substrate in an organic solvent using a hydrophobic sugar derivative. The case using a sugar derivative is introduced here.

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

45

O R

CH2 OH CHOH O OH

+

O R

OH

O O

Sugar acetal

Lipase/non-aqueous medium

H2O

Fatty acid

O CH2 CHOH O OH

O

R O

H+/H2O

HO HO

O OH OH

O O

Sugar acetal ester

Sugar fatty acid ester

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Figure 7. Synthesis of acyl saccharide by condensation of sugar acetal with fatty acid.

There are several hydrophobic sugar derivatives which can become the raw materials of acyl saccharides. For example, hydrophobic derivatives of glucose include isopropylidene glucose (1,2-O-isopropylidene--D-glucofuranose) which is one of the sugar acetals, alkyl glucosides such as ethyl glucoside, and phenylboronic acid ester of glucose. Among them, using the isopropylidene derivative is discussed because the derivative can be easily synthesized and needs relatively low toxic reagents for the synthesis. If a sugar has a structure of cis-diol in its molecule, it reacts with acetone under the presence of an acid catalyst, and a hydrophobic derivative having a dioxolane ring is easily formed. This is usually used as a step for the introduction of a protecting group. At the same time, the solubility in an organic solvent increases significantly. For example, solubility of glucose is only 0.44 mmol/L in acetone (22C) [18], and 0.093 mmol/L in acetonitrile (50C) [28]. Meanwhile, the solubility of isopropylidene glucose is more than 200 mmol/kg at 40C and is extremely high [36]. In addition, diisopropylidene galactose consisting of 1 molecule of galactose and 2 molecules of acetone can be mixed at an arbitrary content with an organic solvent such as 2-methyl-2propanol. If a primary hydroxyl group of the isopropylidene derivative is free, the derivative can become a substrate for the lipase-catalyzed condensation. After dissolving isopropylidene derivative and fatty acid in an organic solvent at high concentrations, a precursor of the acyl saccharide can be synthesized by the addition of lipase (Fig. 7). The precursor is easily converted into an acyl saccharide by deprotection of an isopropylidene group under the presence of an acid catalyst. There are some reports in which the sugar derivative was converted into a corresponding ester efficiently. Fatty acids such as octanoic acid and myristic acid, palmitic acid were used as an acyl substrate for the condensation, and isopropylidene glucose and diisopropylidene galactose, maltose triacetal etc. were used as a sugar derivative [36-38]. When isopropylidene glucose and palmitic acid were condensed by C. antarctica lipase, a corresponding ester was synthesized [36]. The following phenomenon was observed during this reaction. When two substrates reacted in 2-methyl-2-propanol, the reaction rate was high, but the yield on 80 h was low. Meanwhile, the yield on 80 h was high in acetone although the reaction was slow. Both were improved when the mixture of 2-methyl-2-propanol and acetone was used at the volumetric ratio of 25/75 as a reaction medium. Although 2-methyl2-propanol reduces the equilibrium constant as mentioned above, it is very effective for improvement of the reaction rate. On the other hand, an extreme fall of the reaction rate was observed when isopropylidene glucose was used at high concentration for the condensation

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

with palmitic acid. The same phenomenon was also observed when isopropylidene glucose and octanoic acid were condensed [37]. Although the details for this phenomenon are unclear at present, the substrate inhibition may occur at high substrate concentration.

3. KINETICS FOR THE LIPASE-CATALYZED CONDENSATION

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3.1. Reactivity of Substrates for the Lipase-Catalyzed Condensation In the previous section, reaction equilibrium for the synthesis of esters was discussed. Meanwhile, there is another important factor for the lipase-catalyzed reaction; it is the reaction rate. The product can not be obtained virtually when the reaction rate is extremely small even if the equilibrium constant is great. Therefore it is also important to understand the kinetics for the lipase-catalyzed condensation. In this section, quantitative analysis for the substrate specificity of C. antarctica lipase is introduced from the kinetic viewpoint when a carboxylic acid with a short acyl chain was used. Not only a fatty acid but also a carboxylic acid with a short acyl chain can become the substrate of the lipase-catalyzed condensation. Carboxylic acid and alcohol are condensed by C. antarctica lipase to form the corresponding ester. In the condensation, there is a case that the reaction rate is extremely small [39]. The factors for the drop of the reaction rate were quantitatively related to the properties of carboxylic acids [40]. Various kinds of short chain carboxylic acids were condensed with p-methoxyphenethyl alcohol by C. antarctica lipase in acetonitrile, and its reaction rate was measured. Then the maximum reaction rate, Vmax, and the Michaelis constant, Km, of the lipase were estimated. The reaction rate largely depended on the kind of a carboxylic acid. When an alkyl group of a carboxylic acid had a branched structure, in other words, in the case such as isobutyric acid and cyclohexanecarboxylic acid, the reaction rate was very low. The V/Km value was approximately one order lower than that of a carboxylic acid without a branched chain. In addition, the Km value was great when a branched-chain existed. To quantitatively relate the molecular structure of a carboxylic acid to the Km value, the projection area of the alkyl group of a carboxylic acid molecule was estimated, where it was supposed that each atom size in a molecule was van der Waals radius. The Km values were plotted against the estimated projection areas. The plot gave the straight line which was upward toward the right (Fig. 8). This result indicates that, with increasing the projection area, the Km value becomes greater and reactivity of the substrate lowers. Acrylic acid has a double bond conjugated with a C=O bond of the carboxyl group. Therefore, a state of the carboxyl group may be different from the case without the conjugation. Therefore an attention was next paid to the presence of a conjugation in the molecule. The electron density of a carboxyl carbon of carboxylic acids, such as propionic acid and acrylic acid, was estimated as a property of the conjugated double bond. When a conjugated double bond was present, the electron density was low and the carboxylic acid was shown to be hardly attacked by lipase. As a result, the V/Km value decreased approximately one order. Although the electron density was much lower for benzoic acid which has an aromatic ring, its V/Km value was almost at the same level as other conjugated carboxylic acids.

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47

10-2

V/Km [1/min]

A

5 3

10-3

4 1

10

2

8

10-4

9 6

7

10-5

Km [mmol/L]

120

B

80

40

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0 14

18

22

26

30

Projection area [Å2] Figure 8. Effects of projection area of non-carboxylic region of carboxylic acid on (A) V/Km and (B) Km values. Labels represent the carboxylic acid: 1, acrylic acid; 2, crotonic acid; 3, vinylacetic acid; 4, propionic acid; 5, butyric acid; 6, benzoic acid; 7, methacrylic acid; 8, isobutyric acid; 9, isovaleric acid; 10, cyclohexanecarboxylic acid.

As mentioned, the nearby structure of a carboxyl group influences greatly on the reactivity of the substrate for the lipase-catalyzed synthesis of esters. The V/Km value decreases about one order of magnitude by the presence of a branched structure or a conjugated double bond, and the presence of both structures lowers the V/Km value by about two orders of magnitude. In addition, two straight lines are almost parallel in the semilogarithmic plot of the V/Km value against the projection area as shown in Fig. 8. This fact indicates that the presences of a conjugated double bond and a branched structure independently lower the V/Km value.

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

3.2. Kinetics of the Lipase-Catalyzed Condensation of Fatty Acid and Alkyl Glucoside Synthesis of the acyl saccharide by lipase has been widely performed. However, the kinetic analysis for the condensation reaction has not been performed very much. One of the reasons is the low solubility of a sugar in an organic solvent, and it is difficult to estimate the exact kinetic parameters. Therefore octyl glucoside, which was a hydrophobic derivative of glucose and is soluble in an organic solvent, was used instead of an unmodified sugar. Octanoic acid and octyl glucoside was condensed in acetonitrile using C. antarctica lipase, and the kinetic analysis was performed under various reaction conditions based on the ping pong Bi Bi mechanism [41]. The condensation was performed under various conditions. At the low water concentration ( 175 min., the conversion leveled off (ca. 70%). The product concentration of abut 18 mmol/L was accomplished for every kind of the ester. The concentration of the product was equivalent to ca. 6 g/L when lauric acid was used as a substrate. This value was not very high but was not extremely low. Therefore, it is possible to realize the continuous production. The mean residence time was set at 175 min., and the long-term operation was evaluated with changing the kind of fatty acids. The CSTR could be operated stably for at least 11 days as shown in Fig. 12. When lauric acid was used as a substrate, the productivity at ca. 60 kg/(m3·d) was maintained. In addition, remarkable inactivation of the immobilized lipase was not observed during the operation.

4.3. Semi-Continuous Production of Kojic Acid Ester Kojic acid is a substance that has biological functions, such as an antimicrobial activity, a metal-chelating activity, and a tyrosinase inhibitory activity. However, its solubility in oil is low. To raise the solubility in oil, lipase-catalyzed condensation of kojic acid and fatty acid had been carried out in a small-scale batch reactor [53]. A fatty acid remains unreacted even if it is assumed that the conversion is 100% because an excess amount of fatty acid is usually used in the reaction. This is reasonable to improve the yield from the viewpoint of

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

equilibrium, but it is necessary to consider the efficient recovery of the product and the reuse of unreacted substrates for the industrialization. Therefore the system was constructed, in which lauroyl kojic acid was synthesized by a semi-continuous process using a CSTR, and unreacted substrates were reused [54]. The process was proposed as schematically shown in Fig. 13. Kojic acid (40 mmol/L) and the excess amount of lauric acid (100 mmol/L) were mixed in acetonitrile, and it was pumped to the CSTR, the working volume of which was 200 mL. Lipase-catalyzed condensation was performed at 50C in the CSTR (mean residence time = 10 min.), and undissolved kojic acid was recovered after the reaction by filtration at room temperature. The dependences of the solubility of kojic acid, lauric acid, and lauroyl kojic acid on temperature were different in acetonitrile. At lower temperature, the solubilities were in the order of lauric acid > kojic acid >> lauroyl kojic acid (product), and the solubility of the product was extremely lower than that of the other substrates. To recover the product by taking advantage of the difference of the solubility, the reaction mixture was cooled down. In the cooled solution, most of the product and kojic acid precipitated, and the precipitate were recovered efficiently by filtration. Meanwhile, unreacted lauric acid remained dissolved in the solution. Therefore this solution was reused for the next reaction-cycle after dehydration. Because part of kojic acid was precipitated with the product after cooling down, both were separated by washing the precipitate with water. Unreacted kojic acid was used again for a further reaction after dehydration. As a result of this semi-continuous operation, 37 g of the product was obtained after 4 times of the operations. This result corresponds to 53% conversion. Further repetition of the operation will improve both the conversion and the recycle rates of the substrates.

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Filtration

(ppt)

Reactor (CSTR)

Dehydration

Kojic acid Lauric acid

(sup)

Kojic acid (sup)

(sup) (ppt)

Cooling

Filtration

Washing (ppt) Lauroyl kojic acid (Final product)

Figure 13. Semi-continuous system for the production of lauroyl kojic acid using the CSTR. The abbreviations sup and ppt represent supernatant and precipitate, respectively.

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

55

By the combination of the ester synthesis using a CSTR, the recovery of the product by cooling, and the reuse of the unreacted substrates, lauroyl kojic acid could be synthesized efficiently in large quantities. This semi-continuous production system will be applicable to the reaction that has a similar property and may contribute to the continuous synthesis of esters by an immobilized lipase.

5. FUNCTIONALITIES OF THE PRODUCTS 5.1. Surfactant Properties of Acyl Saccharides, Ascorbates and Sugar Alcohols The surface tensions of the aqueous solutions of the acyl mannose [51], ascorbates [50, 55] and sugar alcohols [56] synthesized through the lipase-catalyzed condensation in an organic solvent were measured at the various concentrations by the Wilhelmy method. The critical micelle concentration, CMC, was estimated from the intersection of the two lines in the plots for the dependence of the surface tension on the concentration of the produced esters. The surface excess, , was evaluated from the slope of the line drawn at the low concentrations according to the following equation:

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

d RT   d log C 0.434

(8)

where  is the surface tension, C is the concentration of the product, R is the gas constant, and T is the absolute temperature. The reciprocal of the  value gives the residual area per molecule, a. The CMCs of octanoyl, decanoyl, lauroyl and myristoyl mannoses at 25oC are calculated. The CMC of an acyl mannose was the almost same as that at 37oC of the monoacyl fructoses, which were estimated from Fig. 7 in the literature [57], with the same acyl chain, though the temperature was different. The change in the CMC as a function of the chain length, nCL, is expressed by [58] log CMC  

w nCL  b kT

(9)

where w is the cohesive energy change per methylene group passing the bulk of the solution to the micelle, k is Boltzmann‘s constant, and b is a constant. The w values were estimated to be 2.0 × 10–21 and 1.6 × 10–21 J for the 6-O-acyl mannoses and fructoses, respectively. These values were somewhat smaller than those for the 1-O-alkyl glycosides (glucosides, galactosides, and fucosides: 2.2  2.7 × 10–21 J) [59]. The a values were practically the same among the acyl mannoses tested, and were ca. 0.40 nm2. The a values of the alkyl glycosides were also in the range of 0.37 to 0.49 nm2 [59]. Since acyl mannose molecules would be oriented so as to stick their acyl residues into the air, the a value seemed to be exclusively determined by the saccharide moiety. The situation would be the same for the alkyl glycosides. These would be the reasons why the acyl mannoses and alkyl glycosides had similar a values.

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

Surface tension [mN/m]

80

(a)

(b)

(c)

(d)

60

40

20 10-6

10-4 10-2 10-4 10-2 10-4 10-2 10-4 Concentration of acyl ascorbate [mol/L]

10-2

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Figure 14. Surface tensions of (■) decanoyl and (○) lauroyl ascorbates in distilled water at (a) 30, (b) 35, (c) 40 and (d) 45oC.

The CMCs of acyl ascorbates (C6 12) with longer acyl chains were lower at 25oC, while the  values scarcely depended on the acyl chain length. The CMC and a values of the acyl ascorbates were also estimated. The w values for the acyl ascorbates were calculated from Eq. (9) to be 1.3 × 10–21 J and were much smaller than those for the acyl mannoses and alkyl glucosides. The a values of the acyl ascorbates were ca. 0.30 nm2, which was much smaller than those of the acyl mannoses and alkyl glucosides. The molar volumes of mannose and glucose were 0.114 L/mol [60]. If the hexoses are assumed to be spherical, the crosssectional area is 0.40 nm2, which coincides with the a value evaluated from the surface tension measurement. The molar volume of ascorbic acid was estimated to be 0.106 L/mol, from which the cross-sectional area was calculated to be 0.38 nm2 under the assumption that the molecule was a sphere. The a value of 0.30 nm2 was much smaller than the crosssectional area. Ascorbic acid has a -lactone ring, which is smaller than the pyranose one of mannose or glucose. The -lactone ring might act as the hydrophilic moiety of the acyl ascorbate. Furthermore, the surface tensions of the decanoyl and lauroyl ascorbates dissolved in distilled water were measured at various temperatures by the Wilhelmy method to determine the temperature dependence of the CMC values (Fig. 14). The CMC and a values of each ascorbate are plotted versus the 1000/T values in Fig. 15. The temperature dependence of the CMC was very weak for both ascorbates. The a value was also independent of the temperature, and was ca. 0.35 nm2 for both the decanoyl and lauroyl ascorbates. The effect of pH on the surface tensions of the decanoyl ascorbate solution at 30oC was examined. The CMC value was higher at the higher pH (Fig. 15). The pKa value of L-ascorbic acid is 3.77 [61]. If we assume the pKa value of the ascorbyl moiety is the same as that of ascorbic acid, the degree of dissociation of the ascorbyl moiety increases at the higher pH. The exponential increase in the CMC with an increase in pH could be explained by the increase in the degree of dissociation [62]. The a value was almost constant at any pH, and the value was the same as those obtained at different temperatures for the decanoyl and lauroyl ascorbates.

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

57

1000/T [K-1] 3.2

3.3

3.4 60

-3.0

40

-3.5

20

a [nm2]

log CMC(mol/L)

3.1 -2.5

0

-4.0 1

2

3

4 pH

5

6

7

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Figure 15. Effects of pH and temperature on the critical micelle concentration, CMC, and the residual area per molecule, a, of the decanoyl (closed symbol) and (open symbol) lauroyl ascorbates.

Each of the 1-O-monoacyl sugar alcohols was synthesized in acetone through the immobilized-lipase-catalyzed condensation using erythritol, arabitol, ribitol, xylitol or sorbitol, and octanoic, decanoic, lauric or myristic acid. The surfactant properties of the monoacyl sugar alcohols at 25oC were mostly influenced by the acyl chain length. The CMC value was lower for the sugar alcohol ester with the longer acyl chain. The logarithms of the CMC values were proportional to the hydrophilic-lipophilic balance, HLB, values for each of glycerol, erythritol and xylitol esters. The HLB is an empirical but useful quantity for the practical use of surfactants. Surfactants with the low HLB values of 3 to 8 are, in general, better suited for the preparation of water-in-oil emulsions, whereas the surfactants with the high HLB values of 9 to 18 are suitable for the preparation of oil-in-water emulsions. Although there are some definitions for the HLB value, we calculated the HLB values for the tested monoacyl sugar alcohols according to the Griffin equation.

M  HLB  20 H   M 

(10)

where MH and M are the masses of the hydrophilic moiety and whole surfactant molecule, respectively. The a values of the tested monoacyl sugar alcohols were almost the same at all temperatures, and ranged from 0.3 to 0.4 nm2. They were similar to the value of lauroyl glucose (0.37 nm2), and were smaller than those of lauroyl sucrose (0.56 nm2), raffinose (0.67 nm2) and stachyose (0.72 nm2) [63]. The a value was smaller for the ester with the smaller hydrophilic moiety, and increased as the carbon number of the moiety increased. The crosssectional area of a single alkyl tail has been reported to be 0.22 nm2 [64]. These results also indicated that the a value was exclusively determined by the size of the hydrophilic moiety, and that the monoacyl sugar alcohol would be oriented so as to stick its acyl residue into the

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

air. The critical packing parameter, CPP, is a parameter to express the self-organized structure, and is defined by the following equation [65]:

CPP 

v a0lc

(11)

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where v is the average volume occupied by an alkyl chain with n carbon atoms, a0 is the optimal head group area and lc is the average length of an alkyl chain. The v and lc values in units of nm2 and nm were estimated by Eqs. (12) and (13), respectively, according to the equations proposed by Tanford [66].

v  0.0274  0.0269n

(12)

lc  0.15  0.1265n

(13)

The a value can be used instead of the a0 value. The CPP values of all the monoacyl sugar alcohols were between 0.5 and 1.0 for the monoacyl sugar alcohols except for monooctanoyl xylitol, indicating that the monoacyl sugar alcohols are expected to aggregate in the shape of a bilayer. The value of the monooctanoyl xylitol was 0.43, and it was thought to aggregate into a rod-like form. Although the size or the number of hydroxyl groups of the hydrophilic moieties was different among the esters, the w value in Eq. (9) was common for all the esters and was 2.3 × 10–21 J. The surface tension curves were almost the same for the lauroyl esters of arabitol, ribitol and xylitol, which are isomers of each other. This fact indicated that the orientation of the hydroxyl groups of the sugar alcohols seemed to not significantly affect their surfactant properties. For some of the monoacyl sugar alcohols, the surface tensions were measured at 25oC and 39oC. No significant effect of the temperature on the surface tension was recognized for the esters. The free energy of micellization, Gm, is given by

Gm  RT ln(CMC)

(14)

The enthalpy of micelle formation, Hm, can be calculated from the temperature dependence of the CMC value. d log CMC H m  d (1 / T ) 2.30 R

(15)

The CMC values were measured at 25, 32 and 39oC for the xylitol esters with different acyl chains and for monolauroyl arabitol, ribitol and sorbitol, and there was no significant temperature dependence of the CMC for all the esters. This indicated that the Hm value was almost zero. Therefore, the Hm values for all the tested monoacyl sugar alcohols were assumed in order to estimate the entropy of micellization, Sm, by the following equation:

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase …

S m 

H m  Gm T

59 (16)

The Sm values estimated at 25oC were positive for all the monoacyl sugar alcohols, and increased with the increasing acyl chain length. The positive value seems to be ascribed to the disorder or rupture of the iceberg structure of water by the incorporation of the hydrophobic acyl chain into the structure [67].

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5.2. Bacteriostatic Activities of Acyl Saccharides and Sugar Alcohols Generally, many surfactants have an antimicrobial ability. Acyl glycerols and sucroses, which are also called sugar esters, are commonly used to prevent or suppress bacterial spore development in canned soft drinks stored at 50 to 70oC in the cold and cool seasons. Therefore, the antimicrobial activities of myristoyl, palmitoyl, or stearoyl hexoses, which were glucose, mannose, and galactose, against three Gram-positive bacteria, Bacillus coagulans, Bacillus subtilis and Bacillus licheniformis, were measured in order to investigate the availability of monoacyl hexose as an antimicrobial co-agent [68]. Ten grams of peptone, 2 g of yeast extract and 1 g of magnesium sulfate heptahydrate were added to distilled water, and 1 L of the liquid medium was prepared. The pH of the mixture was adjusted to 7.0 using 1.0 mol/L sodium hydroxide. The medium was autoclaved at 121oC for 20 min. A freezedried type culture of B. coagulans was rehydrated with the medium, and the inoculum was incubated by slowly shaking in a water-bath for 18 h under anaerobic condition at 37oC. The optical density, which was abbreviated OD, at 600 nm of the culture was measured using a spectrophotometer, and then the culture was diluted by the medium to adjust the initial OD of the next generation culture to ca. 0.1. The culture was incubated at 37oC under anaerobic condition. At appropriate intervals, the OD of the sampled culture was measured at 600 nm. The OD of the culture without monoacyl hexose was evaluated as the control (ODcontrol). The ratio of the OD of the culture with monoacyl hexose to ODcontrol was used as an index for the antimicrobial activity. Figure 16 shows the growth inhibition of B. coagulans by monoacyl glucose, mannose, and galactose with various acyl chain lengths from 8 to 18 at the concentration of 40 mg/L. The ratio of the OD for the culture with octanoyl, decanoyl, lauroyl, or myristoyl glucose to the ODcontrol did not change with time, indicating that these esters could not inhibit the growth at this concentration. The palmitoyl and stearoyl glucoses, however, exhibited an antimicrobial activity after 15 h. The myristoyl and stearoyl mannoses also inhibited the growth, though these esters gradually suppressed the growth from the beginning. Myristoyl galactose slowly inhibited the growth, but the antimicrobial activity was low. The OD/ODcontrol value for the culture with palmitoyl or stearoyl galactose more rapidly decreased than that with the myristoyl ester. Glucose esters with acyl chain lengths from 8 to 12 hardly exhibited the antimicrobial activity at 24 h. The myristoyl ester exhibited a very low antimicrobial activity for each hexose, whereas the palmitoyl or stearoyl ester more strongly inhibited the growth. Although the inhibition processes were somewhat different among the monoacyl hexoses, the antimicrobial activities of the monoacyl hexoses with an acyl chain length from 14 to 18 at 24 h were the same.

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

1.0

0.8 (a)

OD/OD control

0.6 1.0

0.8 (b) 0.6 1.0

0.8 (c) 0.6

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0

10 Time [h]

20

Figure 16. The growth inhibition of Bacillus coagulans by (a) glucose, (b) mannose, and (c) galactose esters condensed with (○) octanoic, (□) decanoic, (△) lauric, (●) myristic, (■) palmitic, or (▲) stearic acid at the concentration of 40 mg/L and 37oC.

The bacteriostatic abilities of monoacyl sugar alcohols with different acyl chains and hydrophilic heads were examined against some thermophilic sporeformers, which were the Gram-positive bacteria included Geobacillus stearothermophilus, Bacillus coagulans, Bacillus cereus, Bacillus subtilis, Moorella thermoacetica and Alicyclobacillus acidocaldarius [69]. For comparison, Gram-negative Saccharomyces cerevisiae and Eschericia coli were also used. A monoacyl sugar alcohol was dissolved in water at 1250 mg/L by heating in a microwave oven, and then added to an agar medium to give a specified concentration. The concentration was in the range of 1.6 to 100 mg/L. The medium was autoclaved at 121oC for 15 min. Bacterial spores, which had been heat-shocked at 80oC for 10 min, were inoculated on the agar plate at ca. 104 to 106 CFU for the Gram-positive bacteria, and then cultivated at a temperature adequate for the growth of each bacterium. The pH of the incubation solution for A. acidocaldarius was adjusted to 4.0 by adding 10% (w/v) tartaric acid. After a 7-day incubation, the number of colonies was counted to determine the minimum inhibitory concentration of each monoacyl sugar alcohol for every microorganism. The minimum inhibitory concentrations, MIC, of the monoacyl erythritols and xylitols with

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different acyl chains versus were evaluated. There was a tendency that the monomyristoyl esters had strong bacteriostatic abilities against the thermophilic sporeformers regardless of the hydrophilic head type. The monomyristoyl erythritol and xylitol had the same abilities as the monomyristoyl diglycerol or higher for suppressing the germination of the thermophilic bacteria. Especially, monomyristoyl xylitol seemed to be the most efficacious bacteriostatic agent for the thermophilic Gram-positive bacteria among the tested esters. All the esters lacked the ability to suppress the Gram-negative yeast and bacterium. The bacteriostatic abilities of the various monomyristoyl sugar alcohols against three thermophilic Grampositive bacteria and yeast were measured. The sugar alcohol type significantly affected their ability. Although arabitol, ribitol and xylitol are sugar alcohols with 5 carbons and only the orientation of the hydroxyl group is different, their bacteriostatic abilities were different, especially against B. cereus. All the tested esters were very effective in suppressing the germination of A. acidocaldarius, which is a strictly aerobic bacterium and is often a contaminate in fruit juice. Thus, both the number and orientation of the hydroxyl group of the monoacyl sugar alcohols play important roles in their bacteriostatic abilities as well as the acyl chain length.

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5.3. Antioxidative Properties of Acyl Ascorbates and Alkyl Ferulates Much attention has been paid to the use of polyunsaturated fatty acids, PUFAs, as components in foods [70], but PUFAs are susceptible to autoxidation [71] and causes deterioration of the foods. Some PUFA ascorbates were synthesized using an immobilized lipase from C. antarctica, and their oxidation processes were then observed [72-74]. Linoleic, -linolenic, -linolenic, arachidonic, eicosapentaenoic, docosahexaenoic and conjugated linoleic acids were used. The effect of the molar ratio of the unmodified ascorbic acid or linoleoyl ascorbate to linoleic acid on the suppression of the oxidation of linoleic acid was examined. About 11 to 20 mg of PUFA ascorbate was dissolved in 1 mL of methanol. The solution (20 L) was placed in flat-bottomed glass cups (1.5 cm i.d. and 3.0 cm height), and the methanol was then evaporated under reduced pressure. The cups were placed in a desiccator with a Petri dish containing phosphorus pentoxide to regulate the relative humidity at nearly 0%. The desiccator was stored in the dark at 65oC. At appropriate intervals, 10 L of methyl palmitate solution dissolved in hexane at a concentration of 0.025 mL/mL-hexane was added to the cup as the internal standard for the GC analysis. After the evaporation of hexane, an aliquot (200 L) of sodium methoxide, dissolved in methanol at a concentration of 1.0 g/L, was added. The cup was stored at 70oC for 30 min to transesterify the PUFA ascorbate to the PUFA methyl ester. The methanol was removed under reduced pressure, and the remainder was used for the GC analysis with an FID [75]. Figure 17 shows the oxidation processes of the unmodified PUFA and the PUFA moiety of the PUFA ascorbates at 65oC and nearly 0% relative humidity. The unmodified PUFAs were almost completely oxidized within 5 h, whereas all the PUFA ascorbates were significantly resistant to oxidation. The PUFA moiety of 90% or more remained in the unoxidized state during the test period for every ascorbate. The oxidation processes at 65oC and ca. 0% relative humidity of linoleic acid mixed with ascorbic acid at various molar ratios were measured. At the molar ratios greater than or equal to 0.2, the oxidation of linoleic acid was suppressed at least for 20 h even under the severe

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Fraction of unoxidized acyl moiety and fatty acid

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conditions. Linoleoyl ascorbate was mixed with linoleic acid at various molar ratios and the oxidation process of linoleic acid was measured at 65oC and ca. 0% relative humidity. Similar to the case where the unmodified ascorbic acid was added, the oxidation of linoleic acid was almost completely suppressed at the molar ratios of ≧ 0.2. Antioxidative properties of saturated acyl ascorbates in the bulk [76], the oil-in-water type emulsion [77], and the microcapsule systems [47] were analyzed. The dependence on the acyl chain length of the ascorbate was observed in each system. The antioxidant activity of acyl ascorbates with acyl chain lengths of 10 to 18 was measured in an intestinal epithelial cell line, Caco-2 [78]. Figure 18hows the oxidation index of the Caco-2 cells which were treated with ascorbic acid or acyl ascorbate from the lumen side and oxidized by FeSO4. The lower ratio (index) means the higher antioxidant activity. All the acyl ascorbates exhibited the higher activity for the oxidation than unmodified ascorbic acid. Especially, lauroyl and myristoyl ascorbates were the most effective antioxidant among the tested asorbates. An alkyl ferulate is more hydrophobic than ferulic acid. Solubilities of 1-pentyl ferulate, 1-hexyl ferulate, 1-heptyl ferulate, and ferulic acid in linoleic acid were measured at 37–80C [79]. The solubility of alkyl ferulate and ferulic acid rose with increasing temperature. Furthermore, the solubility of each alkyl ferulate was approximately 20 times higher than that of unmodified ferulic acid. However, between three kinds of alkyl ferulates, the remarkable difference was not observed for the solubility in linoleic acid. In addition, the partition coefficients of an alkyl ferulate and ferulic acid to linoleic acid in water/linoleic acid system were also measured [80]. The partition coefficient of the alkyl ferulate (ca. 1150) was extremely greater than that of ferulic acid (ca. 5.1), and most of the alkyl ferulate existed in the oil phase.

1.0

(a)

0.8

(b)

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0.4

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0 0

2

4

0 Time [h]

2

4

6

Figure 17. Oxidation processes of polyunsaturated fatty acids and their ascorbates at 65oC and nearly 0% relative humidity. (a) n-6 series: () -linolenic acid, () -linolenoyl ascorbate, () dihomo-linolenic acid, () dihomo--linolenoyl ascorbate, () arachidonic acid and () arachidonoyl ascorbate; (b) n-3 series; () -linolenic acid, () -linolenoyl ascorbate, () eicosapentaenoic acid, () eicosapentaenoyl ascorbate, () docosahexaenoic acid and () docosahexaenoyl ascorbate.

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Intensity ratio (Oxidation index)

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0.2

0.1

0 N.C.

P.C.

Free AsA

C10

C12 C14 C16 Acyl ascorbates

C18

Fraction of unoxidized linoleic acid

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Figure 18. Antioxidant activity of acyl ascorbates in Caco-2 cells. The N.C. refers to the negative control without any antioxidant, and P.C. indicates the positive control with FeSO4 and without any antioxidant. Free AsA means the unmodified ascorbic acid, and C10 to C18 indicate the carbon number of acyl chains of ascorbates.

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0 0

20

40

60

80

Time [h] Figure 19. Oxidation processes of linoleic acid with (○) ferulic acid, (△) 1-pentyl ferulate, (□) 1-hexyl ferulate, (▽) 1-heptyl ferulate, and (●) without any additive at 37C. The molar ratio of the additive to linoleic acid was 3.0 × 10–4.

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A

Fraction of unoxidized linoleic acid

1.0

0.5

0

B

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0

5

10

15

Time [d] Figure 20. Oxidation processes at 37C of linoleic acid mixed with ( ) 1-hexyl ferulate and ( ) ferulic acid at the molar ratio (additive/linoleic acid) of 1.0 × 10–2, ( ) without any additive, and encapsulated with (A) maltodextrin and (B) gum arabic.

Because ferulic acid is a substance having antioxidative activity, an alkyl ferulate may also have the activity to suppress the oxidation of hydrophobic compounds. The suppressive effects of 1-pentyl ferulate, 1-hexyl ferulate, 1-heptyl ferulate and unmodified ferulic acid toward the autoxidation of linoleic acid were evaluated in a bulk system [79]. Both an alkyl ferulate and ferulic acid exhibited suppressive effect on the oxidation, but an alkyl ferulate more greatly suppressed the oxidation (Fig. 19). The suppressive effect was due to the high solubility of an alkyl ferulate in linoleic acid. The induction period for the oxidation of linoleic acid was elongated, and the rate constant for the oxidation also tend to become smaller. But no remarkable difference between three alkyl ferulates was observed. In the bulk oxidation system, the suppressive effect of an alkyl ferulate was remarkable. Meanwhile, the suppressive effect was also evaluated in the microencapsulated system [80]. An alkyl ferulate or ferulic acid was added in linoleic acid, and the mixture was encapsulated using maltodextrin or gum arabic as a wall material. Irrespective of the kind of an additive, the oxidation of linoleic acid was suppressed (Fig. 20). The suppression of the oxidation was

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particularly great for gum arabic. One of the reasons seems that gum arabic itself has suppressive effect toward the oxidation. Meanwhile, the oxidation was suppressed to some extent when maltodextrin was used as a wall material under presence of ferulic acid. When an alkyl ferulate was added instead of ferulic acid, the oxidation was more greatly suppressed. This is due to the fact that maltodextrin did not exhibit suppressive effect on the oxidation, and the suppressive effect of an alkyl ferulate appeared conspicuously. On the whole, it was found that alkyl ferulates were well miscible with linoleic acid and effectively suppressed the oxidation of linoleic acid both in the bulk system and microencapsulated system. The addition of an alkyl ferulate contributes both to the elongation of the induction period and the decrease of the rate constant for the oxidation.

5.4. Oxidation of Linoleoyl Residue of its Trehalose Ester in a Micelle System

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The oxidative stability of lipids would be affected by the states in which they exist, such as bulk, emulsion and micelle systems. Linoleic acid was regiospecifically condensed with trehalose using the immobilized lipase in a mixture of 2-methyl-2-propanol and pyridine (6/4 (v/v)) to produce the 6-O-monolinoleoyl trehalose, and the autoxidation of the linoleoyl residue was measured in the micelle of the ester [81]. The oxidation process of the residue could be expressed by the rate equation of the autocatalytic type, and the rate constant was higher for the higher linoleoyl trehalose concentration. However, the constant was not proportional to the concentration at the concentrations higher than its critical micelle concentration, suggesting that the migration of oxygen into the hydrophobic core of the linoleoyl residues through the hydrophilic trehalose layer of the micelles affected the overall oxidation rate.

5.5. Stability of Acyl Ascorbates 6-O-Monoacyl ascorbates were synthesized by the condensation of ascorbic acid with octanoic, decanoic, lauric, myristic or palmitic acid using immobilized lipase in acetone, and the decomposition processes of the acyl ascorbates in air were measured [82]. First, the decomposition processes of octanoyl, decanoyl, lauroyl, myristoyl and palmitoyl ascorbates at 80oC and at 12, 44 and 75% relative humidity were examined. The relative humidity significantly affected the decomposition of the acyl ascorbates, and all the ascorbates decomposed faster at the higher relative humidity. There was a tendency that an ascorbate with a shorter acyl chain decomposed faster at 12 and 44% relative humidity. The decomposition kinetics of the acyl ascorbates was empirically expressed by the Weibull equation, which is flexible and has a potential for describing many deterioration kinetics [83]: Y  exp[ (kt) n ]

(17)

where Y is the fraction of the remaining acyl ascorbate at time t, k is the rate constant, the reverse of which is called the scale parameter, and n is the shape constant. The kinetic parameters, k and n, were evaluated by fitting the experimental results by nonlinear

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

regression. The k values were the higher at the higher relative humidity for every acyl ascorbate. The acyl chain length affected the k value at 44% relative humidity, and the k value was higher for the shorter acyl chain. The n value for the decomposition of any acyl ascorbate at 75% relative humidity was nearly equal to or slightly less than unity, while the n values at 12% and 44% relative humidities were higher than unity. The decomposition processes of the acyl ascorbates at 37, 50 and 65oC and at a 75% relative humidity were also shown. Every acyl ascorbate decomposed faster at the higher temperatures. The acyl-chain length also affected the decomposition of the ascorbates, and the ascorbate having a shorter acyl chain decomposed faster. Although the k value was higher at the higher temperature, there was no significant temperature dependency for the n value and the values were almost 1.0 ± 0.4. The k value for the decomposition of the ascorbate having the shorter acyl chain was higher. The temperature dependence of the rate constant k was analyzed based on the Arrhenius equation: k  k 0 exp( 

E ) RT

(18)

where k0 is the frequency factor, E is the activation energy, R is the gas constant, and T is the absolute temperature. Both the E and k0 values were higher for the ascorbate with the longer acyl chain. The E values are plotted versus the natural logarithms of the k0 values. The plots lie on a straight line (R2 = 0.997). Equation 3 is one of the expressions describing the enthalpy-entropy compensation [84].

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E  RT lnk 0  

(19)

where T is an isokinetic temperature and  is a constant. The linear relationship indicated that the compensation held during the decomposition and that the decomposition of every acyl ascorbate essentially proceeded by the same mechanism. The T value was estimated to be 101oC from the slope. At this temperature, the rate constant k for the decomposition of all the acyl ascorbates would be the same. The decomposition of the acyl ascorbate would consist of the oxidative degradation of the ascorbyl moiety and the hydrolysis of the ester bond between the acyl and ascorbyl moieties. Figure 21 shows the transient changes in the fractions of the remaining lauroyl ascorbate and lauric acid liberated by the hydrolysis of lauroyl ascorbate at 65oC and 75% relative humidity. During the early stage during the storage, the decomposition of lauroyl ascorbate rapidly proceeded, but lauric acid was scarcely formed. The lauric acid was gradually liberated when the fraction of the remaining lauroyl ascorbate became less than 0.5. The relationship between the fractions of the consumed lauroyl ascorbate and liberated lauric acid is shown in the inset of Fig. 21. The amount of consumed lauroyl ascorbate was calculated by subtracting the fraction of the remaining lauroyl ascorbate from unity. The broken line in the inset of Fig. 21 was drawn assuming that the disappearance of the lauroyl ascorbate was ascribed to its hydrolysis. All the plots were under the broken line, thus no lauric acid was liberated during the early stage of the decomposition of the lauroyl ascorbate. This fact indicated that the oxidative degradation of the ascorbyl moiety first occurred and then the hydrolysis of the ester bond followed to liberate the lauric acid. That is, the decomposition of acyl ascorbate would be a consecutive process.

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Fraction of liberated lauric acid

Fractions of remaining lauroyl ascorbate and liberated lauric acid

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0.5 1 Fraction of consumed lauroyl ascorbate

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5

10 15 Time [days]

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25

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Figure 21. Decomposition of () lauroyl ascorbate and formation of () lauric acid at 65ºC and 75% relative humidity. The solid curve was calculated using the estimated kinetic parameters of the Weibull model, and the broken curve was empirically drawn. The inset shows the relationship between the fractions of the consumed lauroyl ascorbate and liberated lauric acid. The solid curve in the inset indicates the formation of lauric acid based on the assumption that the lauroyl ascorbate is first hydrolyzed to lauric acid.

The decomposition of saturated acyl ascorbate in an aqueous solution was measured at various pHs and temperatures [85]. The buffer solutions used were 0.1 mol/L sodium citrateHCl for pHs 2, 3 and 4, 0.1 mol/L sodium citrate-NaOH for pHs 5 and 6, and 0.1 mol/L TrisHCl for pHs 7, 8 and 9. The stability of acyl ascorbates was assessed at 30, 40, 50 and 60oC. The transient changes in decanoyl ascorbate concentration at 40oC and various pHs were measured. The initial ascorbate concentration was 2.0×10–4 mol/L at every pH. Decanoyl ascorbate was the most stable at pH 3 in the tested range. The decrease in the ascorbate concentration was faster at the higher pHs. The ascorbate also decreased faster at pH 2 than at pH 3. The decrease with time at pH 3 showed a clear sigmoidal pattern. This pattern was also recognized at pH 2, 4, 5 and 6, although it was weak. The rate constant k in the Weibull model (Eq. (17)) was the lowest at pH 3, and it increased almost exponentially with an increase in pH. On the other hand, the shape constant n was the largest at pH 3, and there was a tendency that the constant was lower at higher pH. The largest n value at pH 3 reflected the strong sigmoidal pattern. Because the C/C0 values at pHs 7 to 9 simply decreased with time, the n values at those pHs were almost 1 and the degradation of the ascorbate obeyed pseudo first-order kinetics. In order to know which mechanism causes the decrease in the ascorbate concentration, the increase in the decanoic acid concentration was measured at pHs 3, 6 and 9 as well as the decrease in the decanoyl ascorbate concentration. No decanoic acid was

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Yoshiyuki Watanabe, Takashi Kobayashi and Shuji Adachi

detected within 24 h at pH 3 although the half of the decanoyl ascorbate disappeared at that time. A slight amount of decanoic acid was liberated at pH 6. Therefore, the decomposition of the ascorbyl moiety was the predominant mechanism for its disappearance at a pH equal to or lower than 6. At pH 9, the decanoic acid concentration increased during the early stage and its increase continued even after decanoyl ascorbate completely disappeared. The increase in the latter stage would be ascribed to the hydrolysis of the oxidized product of acyl ascorbate such as acyl dehydroascorbate. The initial rate of the decrease in the decanoyl ascorbate concentration was higher than that of the increase in the decanoic acid concentration. Therefore, the decomposition of the ascorbyl moiety and the hydrolysis of the ester bond should simultaneously occur at pH 9. These results indicated that the hydrolysis became more significant at higher pHs although the decomposition of the ascorbyl moiety was the predominant mechanism for the decrease in the decanoyl ascorbate concentration. The degradation of decanoyl ascorbate at pH 5 and 40oC was measured at different initial concentrations. The initial concentration was changed from 5.0×10–5 to 1.0×10–2 mol/L. The kinetic parameters, k and n, in the Weibull model were estimated at the respective initial concentrations, and were plotted versus the initial concentration. There were no significant dependencies of the shape constant, n, and the rate constant, k, on the initial concentration. The degradation of acyl ascorbates was measured at pH 5 and different temperatures. The initial concentration of each ascorbate was fixed at 2.0×10–4 mol/L. Both the E and k0 values in Eq. (18) for the degradation seemed to depend on the acyl chain length of ascorbate. As mentioned above, the hydrolysis of the ester bond did not significantly occur at pH 5, but only the oxidation of the ascorbyl moiety of the ascorbate was responsible for the decomposition of each acyl ascorbate.

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ACKNOWLEDGMENTS Most of this study was supported by the Program for the Promotion of Basic Research Activities for Innovative Biosciences (PROBRAIN), Japan, and it was partly supported by a Grant-in-Aid for Young Scientists (B) 19780106 from the Ministry of Education, Culture, Sports, Science and Technology (MEXT), Japan. The authors are very grateful to Dr. Ryuichi Matsuno, Professor Emeritus of Kyoto University and the President of Ishikawa Prefectural University, for his valuable discussions and advises throughout this study, and sincerely appreciate Dr. Kazuhiro Nakanishi, Professor Emeritus of Okayama University and Professor of Chubu University, for his patient and accurate guidance. The authors would like to thank all the members of the Laboratory of Biomolecular Engineering, Department of Biotechnology and Chemistry, Faculty of Engineering, Kinki University, and those of the Laboratory of Bioengineering, Division of Food Science and Biotechnology, Graduate School of Agriculture, Kyoto University.

REFERENCES [1]

Adachi, S. In Handbook of Biocatalysis; Hou, C. T.; Ed.; Chapter 10 Lipase-catalyzed condensation in an organic solvent; Taylor & Francis: London, 2005; pp 10-1－ 10-15.

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Synthesis of Acyl Saccharides and Ascorbates Using Immobilized Lipase … [2] [3]

[4] [5] [6] [7]

[8]

[9] [10] [11] [12] [13]

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[14] [15] [16] [17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27] [28]

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Adachi, S.; Kobayashi, T. J. Biosci. Bioeng. 2005, 99, 87-94. Adachi, S.; Piao, J. In Biocatalysis and Biotechnology for Functional Foods and Industrial Products; Hou, C. T.; Ed.; Monoacyl Sugar Alcohols Synthesized through Lipase-catalyzed Condensation in an Organic Solvent; Taylor & Francis Books: London, 2006; pp 333-348. Gumel, A. M.; Annuar, M. S. M.; Heidelberg, T.; Chisti, Y. Process Biochem. 2011, 46, 2079-2090. Karmee, S. K. Biofuels, Bioprod. Biorefin. 2008, 2, 144-154. Kobayashi, T.; Adachi, S. Biotechnol. Lett. 2004, 26, 1461-1468. Watanabe, Y.; Adachi, S. In Biochemical Engineering; Dumont, F. E.; Sacco, J. A.; Ed.; Chapter 2 Enzymatic synthesis of acyl ascorbate and its function as a food additive; Nova Science Publishers, Inc.: New York, NY, 2009; pp 41-77. Watanabe, Y.; Adachi, S. In Organic Solvents: Properties, Toxicity, and Industrial Effects; Carter, R. E.; Ed.; Chapter 2 Lipase-catalyzed synthesis of edible surfactants in microaqueous organic solvents; Nova Science Publishers, Inc.: New York, NY, 2011; pp 35-55. Nishikawa, Y.; Yoshimoto, K.; Nishijima, M.; Fukuoka, F.; Ikekawa, T. Chem. Pharm.Bull. 1987, 29, 505-513. Sarney, D. B.; Vulfson, E. V. Trends Biotechnol. 1995, 13, 164-172. Miwa, N.; Yamazaki, H. Exp. Cell Biol. 1986, 245-249. Nagao, N.; Matsubara, H.; Yamanaka, T.; Etoh, T.; Miwa, N.; Iwagaki, H.; Saiki, I.; Yamaoka, S.; Itoh, S.; Kokata, E. Vitamins 1996, 70, 199-200 (in Japanese). Shima, M.; Yohdoh, K.; Yamaguchi, M.; Kimura, Y.; Adachi, S.; Matsuno, R. Biosci. Biotechnol. Biochem. 1997, 61, 1150-1155. Linko, Y. Y.; Hayakawa, K. Trends in Food Sci. Technol. 1996, 7, 59-63. Yoshii, H.; Furuta, T.; Itoh, T.; Yoshimura, K.; Misawa, Y.; Hata, N.; Kobayashi, H.; Linko, Y. Y. Biosci. Biotechnol. Biochem. 1999, 63, 662-665. Adachi, S.; Nagae, K.; Matsuno, R. J. Mol. Catal. B: Enzym. 1999, 6, 21-27. Arcos, J. A.; Bernabe, M.; Otero, C. Enzyme Microb. Technol. 1998, 22, 27-35. Arcos, J. A.; Bernabe, M.; Otero, C. Biotechnol. Bioeng. 1998, 57, 505-509. Degn, P.; Pedersen, L. H.; Duus, J. Ø.; Zimmermann, W. Biotechnol. Lett. 1999, 21, 275-280. Ducret, A.; Giroux, A.; Trani, M.; Lortie, R. Biotechnol. Bioeng. 1995, 48, 214-221. Ljunger, G.; Adlercreutz, P.; Mattiasson, B. Biotechnol. Lett. 1994, 16, 1167-1172. Tsuzuki, W.; Kitamura, Y.; Suzuki, T.; Kobayashi, S. Biotechnol. Bioeng. 1999, 64, 267-271. Cao, L.; Fischer, A.; Bornsceuer, U. T.; Schmid, R. D. Biocatal. Biotransform. 1997, 14, 269-283. Kim, J. E.; Han, J. J.; Yoon, J. H.; Rhee, J. S. Biotechnol. Bioeng. 1998, 57, 121-125. Ducret, A.; Giroux, A.; Trani, M.; Lortie, R. J. Am. Oil Chem. Soc. 1996, 73, 109-113. Watanabe, Y.; Miyawaki, Y.; Adachi, S.; Nakanishi, K.; Matsuno, R. Enzyme Microb. Technol. 2001, 29, 494-498. Khaled, N.; Montet, D.; Farines, M.; Pina, M.; Graille, J. Oléagineux, 1992, 47, 181190. Watanabe, Y.; Miyawaki, Y.; Adachi, S.; Nakanishi, K.; Matsuno R. J. Mol. Catal. B: Enzym. 2000, 10, 241-247.

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[29] Fredenslund, A.; Jones, R. L.; Prausnitz, J. M. AIChE J. 1975, 21, 1086-1099. [30] Reichardt, C. Angew. Chem. Int. Ed. Engl. 1979, 18, 98-110. [31] Asahara, T.; Tokura, N.; Okawara, M.; Kumanotani, J.; Seno, M. Solvent Handbook; Kodansha Scientific: Tokyo, 1976; pp 355-357, 363-365, 507-512, 642-648. [32] Uedaira, H.; Ishimura, M.; Tsuda, S.; Uedaira, H. Bull. Chem. Soc. Jpn. 1989, 63, 33763379. [33] Kobayashi, T.; Furutani, W.; Adachi, S.; Matsuno, R. J. Mol. Catal. B: Enzym. 2003, 24, 61-66. [34] Zhang, X.; Adachi, S.; Watanabe, Y.; Kobayashi, T.; Matsuno, R. Biotechnol. Prog. 2003, 19, 293-297. [35] Kuwabara, K.; Watanabe, Y.; Adachi, S.; Nakanishi, K.; Matsuno, R. Biochem. Eng. J. 2003, 16, 17-22. [36] Kobayashi, T.; Ehara, T.; Mizuoka, T.; Adachi S. Biotechnol. Lett. 2010, 32, 16791684. [37] Kobayashi, T.; Takahashi, T.; Adachi S. J. Oleo Sci., in press. [38] Sarney, D. B.; Kapeller, H.; Fregapane, G.; Vulfson, E. N. J. Am. Oil Chem. Soc. 1994, 71, 711-714. [39] Compton, D. L.; Laszlo, J. A.; Berhow, M. A. J. Am. Oil Chem. Soc. 2000, 77, 513519. [40] Kobayashi, T.; Adachi, S.; Matsuno, R. Biotechnol. Lett. 2003, 25, 3-7. [41] Kobayashi, T.; Adachi, S.; Matsuno, R. Biochem. Eng. J. 2003, 16, 323-328. [42] Yoshida, Y.; Kimura, Y.; Adachi, S. J. Biosci. Bioeng. 2006, 102, 66-68. [43] Kawamura, Y.; Nakanishi, K.; Matsuno, R. Biotechnol. Bioeng. 1981, 23, 488-490. [44] Yoshida, Y.; Kimura, Y.; Kadota, M.; Tsuno, T.; Adachi, S. Biotechnol. Lett. 2006, 28, 1471-1474. [45] Kobayashi, T.; Matsuo, T.; Kimura, Y.; Adachi, S. J. Am. Oil Chem. Soc. 2008, 85, 1041-1044. [46] Kuwabara, K.; Watanabe, Y.; Adachi, S.; Nakanishi, K.; Matsuno, R. Food Chem. 2003, 82, 191-194. [47] Watanabe, Y.; Fang, X.; Minemoto, Y.; Adachi, S.; Matsuno, R. J. Agric. Food Chem. 2002, 50, 3984-3987. [48] Humeau, C.; Girardin, M.; Coulon, D.; Miclo, A. Biotechnol. Lett. 1995, 17, 10911094. [49] Watanabe, Y.; Adachi, S.; Nakanishi, K.; Matsuno, R. Food Sci. Technol. Res. 1999, 5, 188-192. [50] Kuwabara, K.; Watanabe, Y.; Adachi, S.; Nakanishi, K.; Matsuno, R. J. Am. Oil Chem. Soc. 2003, 80, 895-899. [51] Watanabe, Y.; Miyawaki, Y.; Adachi, S.; Nakanishi, K. Biochem. Eng. J. 2001, 8, 213216. [52] Watanabe, Y.; Kuwabara, K.; Adachi, S.; Nakanishi, K.; Matsuno, R. J. Agric. Food Chem. 2003, 51, 4628-4632. [53] Liu, K. J.; Shaw, J. F. J. Am. Oil Chem. Soc. 1998, 75 1507-1511. [54] Kobayashi, T.; Adachi, S.; Nakanishi, K.; Matsuno, R. Biochem. Eng. J. 2001, 9, 85-89. [55] Watanabe, Y.; Adachi, S.; Fujii, T.; Nakanishi, K.; Matsuno, R. Jpn J. Food Eng. 2001, 2, 73-75.

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[56] Piao, J.; Kishi, S.; Adachi, S. Colloids Surf. A: Physicochem. Eng. Aspects 2006, 277, 15-19. [57] Scheckermann, C.; Schlotterbeck, A.; Schmidt, M.; Wray, V.; Lang, S. Enzyme Microb. Technol. 1995, 17, 157-162. [58] Shinoda, K.; Yamaguchi, T.; Hori, R. Bull. Chem. Soc. Jpn 1961, 34, 237-241. [59] Kobayashi, T.; Adachi, S.; Matsuno, R. Biotechnol. Lett. 1999, 21, 105-109. [60] Adachi, S.; Matsuno, R. Biosci. Biotechnol. Biochem. 1997, 61, 1296-1301. [61] Kröger-Ohlsen, M.; Skibsted; J. Agric. Food Chem. 1997, 45, 668-676. [62] Shinoda, K.; Becher, P. Principles of Solution and Solubility; Marcel Dekker: New York, NY, 1978; pp 157-179. [63] Söderberg, I.; Drummond, C. J.; Furlong, D. N.; Godkin, S.; Matthews, B. Colloids Surf. A: Physicochem. Eng. Aspects 1995, 102, 91-97. [64] Kitahara, A.; Tamai, Y.; Hayano, S.; Hara, I. Surfactants: Property, Application and Chemical Ecology; Koudansha: Tokyo, 1979; p 23. [65] Israelachvili, J. N.; Mitchell, D. J.; Ninham, B. W. Biochim. Biophys. Acta 1977, 470, 185-201. [66] Tanford, C. The Hydrophobic Effect: Formation of Micelles and Biological Membranes; 2nd ed.; Wiley: New York, NY, 1980. [67] Frank, H. S.; Evans, M. W. J. Chem. Phys. 1945, 13, 507. [68] Watanabe, Y.; Shirai, Y.; Miyake, M.; Kitano, J.; Adachi, S. Intl. J. Food Prop., in press. [69] Piao, J.; Kawahara-Aoyama, Y.; Inoue, T.; Adachi, S. Biosci. Biotechnol. Biochem. 2006, 70, 263-265. [70] Takahata, K.; Monobe, K.; Tada, M.; Weber, P. C. Biosci. Biotechnol. Biochem. 1998, 62, 2079-2085. [71] Adachi, S.; Ishiguro, T.; Matsuno, R. J. Am. Oil Chem. Soc. 1995, 72, 547-551. [72] Watanabe, Y.; Minemoto, Y.; Adachi, S.; Nakanishi, K.; Shimada, Y.; Matsuno, R. Biotechnol. Lett. 2000, 22, 637-640. [73] Watanabe, Y.; Adachi, S.; Nakanishi, K.; Matsuno, R. J. Am. Oil Chem. Soc. 2001, 78, 823-826. [74] Watanabe, Y.; Sawahara, Y.; Nosaka, H.; Yamanaka, K.; Adachi, S. Biochem. Eng. J. 2008, 40, 368-372. [75] Minemoto, Y.; Ishido, E.; Adachi, S.; Matsuno, R. Food Sci. Technol. Res. 1999, 5, 104-107. [76] Watanabe, Y.; Ishido, E.; Fang, X.; Adachi, S.; Matsuno, R. J. Am. Oil Chem. Soc., 2005, 82, 389-392. [77] Watanabe, Y.; Nakanishi, H.; Goto, N.; Otsuka, K.; Kimura, T.; Adachi, S. J. Am. Oil Chem. Soc. 2010, 87, 1475-1480. [78] Kimura, Y.; Kanatani, H.; Shima, M.; Adachi, S.; Matsuno, R. Biotechnol. Lett. 2003, 25, 1723-1727. [79] Fang, X.; Shima, M.; Kadota, M.; Tsuno, T.; Adachi, S. Biosci. Biotechnol. Biochem. 2006, 70, 457-461. [80] Fang, X.; Kikuchi, S.; Shima, M.; Kadota, M.; Tsuno, T.; Adachi, S. Eur. J. Lipid Sci. Technol. 2006, 108, 97-102. [81] Chen, J.; Kimura, Y.; Adachi, S. Food Sci. Technol. Res. 2006, 12, 163-166.

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[82] Watanabe, Y.; Sawahara, Y.; Asai, S.; Adachi, S. Food Sci. Technol. Res. 2008, 14, 139-143. [83] Cunha, L. M.; Oliveira, F. A. R.; Oliveira, J. C. J. Food Eng. 1998, 37, 175-191. [84] Leffer, J. E. J. Org. Chem. 1955, 20, 1202-1231. [85] Kuwabara, K.; Watanabe, Y.; Adachi, S.; Matsuno, R. J. Food Sci. 2005, 70, 7-11.

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In: Lipase Editors: Hamdi Sassi and Sofien Cannamela

ISBN 978-1-62081-366-9 © 2012 Nova Science Publishers, Inc.

Chapter III

SOLVENT-FREE BIOCATALYTIC SYNTHESIS OF POLYGLYCEROL POLYRICINOLEATE (PGPR) USING IMMOBILISED CANDIDA RUGOSA AND RHIZOPUS ARRHIZUS LIPASES S. Ortega, M. C. Montiel, M. F. Máximo and J. Bastida Chemical Engineering Department. University of Murcia, Campus de Espinardo, Murcia, Spain

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ABSTRACT The use of lipases is continuously increasing due to its ability to catalyse esterification, interesterification, acidolysis, alcoholysis and aminolysis in addition to the hydrolytic activity on triglycerides, to produce industrially important products such as emulsifiers, surfactants, wax esters, chiral molecules, biopolymers, modified fats and oils, structured lipids, and flavour esters. We have developed the biocatalytic synthesis of a food additive named polyglycerol polyricinoleate (PGPR) and identified with the code E-476. PGPR is widely known as an excellent water-in-oil emulsifier in the food industry, because it forms very stable emulsions even when the water content is very high, such as 80%. Therefore, PGPR is used as emulsifier in tin-greasing emulsions for the baking trade, and for the production of low-fat spreads. However, the main application of PGPR is in the chocolate industry, where is used in the adjustment of rheological properties of chocolate, improving the moulding properties of the molten chocolate. An additional property of PGPR in chocolate is its ability to limit fat bloom. The enzymatic synthesis of PGPR by the catalytic action of one or more lipases (which act in mild reaction conditions of temperature and pressure, neutral pH and in a solvent-free system), makes the process environmentally friendly and avoids side reactions so that the obtained product has a higher purity and quality than the current marketed PGPR obtained by chemical processes.

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1. INTRODUCTION Polyglycerol esters have been used as food additives for many years. From the official point of view, food grade polyglycerol esters are divided in two classes: polyglycerol esters of edible fatty acids (E-number: E-475, also known as ―PGFA‖) and polyglycerol polyricinoleate (E-number: E-476, also known as ―PGPR‖). Polyglycerol polyricinoleate is used to maintain stable emulsions of oil and water systems with high water content as well as a viscosity modifier. In the chocolate industry, PGPR is used because it causes a noticeable reduction in the yield stress of molten chocolate. This allows chocolate to be moulded, without any air bubbles, easier coating of particulate ingredients, and the thickness of chocolate coating to be adjusted optimally. An additional property of PGPR in chocolate is its ability to limit fat bloom [1]. Known chemical methods for preparing PGPR involve autocatalytic condensation of ricinoleic acid and alkali-catalysed reaction between the condensed ricinoleic acid and polyglycerol. These procedures have the disadvantage of requiring very long reaction times, involving high energy costs. This fact, together with the high operating temperature can adversely affect the quality of the final product because of problems related with coloration and odours, making it unsuitable for the food industry [2]. As an alternative, the authors in this contribution propose the biotechnological production of PGPR using lipases, which act in mild reaction conditions and produce a final product more suitable for use as a food additive. The enzymatic procedure described in this work consists of two steps. First, the ricinoleic acid is polymerised to obtain the polyricinoleic acid, PR, also known as ricinoleic acid estolide [3-5]. Then, it is esterified with polyglycerol to obtain polyglycerol polyricinoleate, PGPR. Figure 1 shows the reactions involved in the biosynthesis. The optimization of some reaction conditions is especially important in an experimental system like the described one. It is known that temperature is a crucial parameter in every enzyme catalysed reactor but in our case, due to the special characteristics of the reaction medium (solvent-free), temperature greatly influences viscosity, mass transport phenomena and, as a consequence, the esterification rate [8]. While high temperature favours the medium fluidity, enzyme has to be prevented from thermal deactivation [8, 9]. Another decisive parameter in these processes is the water content. Water plays multiple roles in lipase-catalysed esterifications performed in non-conventional media. It is widely known that water is absolutely necessary for the catalytic function of enzymes because it participates, directly or indirectly, in all non-covalent interactions that maintain the conformation of the catalytic site of enzymes [10-12]. On the other hand, in esterification/hydrolysis reactions it is well-known that the water content affects the equilibrium conversion of the reactions. Particularly, in the case of estolides production, the water formed by the reaction must be removed from the reaction mixture if polyricinoleic acid with a high degree of condensation is to be obtained [13]. Usually, enzyme immobilisation improves their operational stability, while preventing contamination of the substrate being converted. In addition, immobilised enzymes can be easily separated from the reaction media for reuse or for use in continuous reactors. For these reasons, efforts have been devoted to obtaining an immobilised derivative with a high immobilised protein percentage and enzymatic activity for the present application [4].

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Figure 1. Biocatalytic synthesis of PGPR. First reaction catalysed by Candida rugosa lipase and second reaction catalysed by Rhizopus arrhizus lipase.

Moreover, because PGPR is a food additive, it should fulfil requests given by the Commission Directive of the European Communities [14], which are summarized in Table 1. In the light of the above, the main objective of this work is the development of the biocatalytic production of PGPR to obtain a higher quality product than the one obtained by chemical processes. Both reactions involved in PGPR obtaining are studied. Different enzymes, free and immobilised, and several reaction equipment are assayed.

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S. Ortega, M. C. Montiel, M. F. Máximo et al. Hence the different objectives of this work are the following: 1) Study of the main operational variables of the polyricinoleic acid production in different reactors with free and immobilised Candida rugosa lipase. 2) Screening of lipases to catalyse the esterification of polyricinoleic acid with polyglycerol. 3) Study of the main operational variables of the esterification of polyricinoleic acid with polyglycerol in different reactors with free and immobilised Rhizopus arrhizus lipase. 4) Selection and study of the immobilisation process of lipases used to catalyse each reaction. Table 1. Purity criteria for E-476, polyglycerol polyricinoleate

Synonyms

Definition Description Identification

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A. Solubility B. Positive tests for glicerol, polyglicerol and for ricinoleic acid C. refractive index [n]65D Purity Polyglycerols Hydroxyl value Acid value Arsenic Lead Mercury Cadmium Heavy metals (as Pb)

Glycerol esters of condensed castor oil fatty acids. Polyglycerol esters of polycondensed fatty acids from castor oil. Polyglycerol esters of interesterified ricinolecic acid. PGPR Polyglycerol polyricinoleate is prepared by the esterification of polyglycerol with condensed castor oil fatty acids. Clear, highly viscous liquid. Insoluble in water and in ethanol. Soluble in ether, hydrocarbons and halogenated hydrocarbons.

Between 1.4630 y 1.4665 The polyglycerol moiety shall be composed of not less than 75% of di-, tri-, and tetraglycerols and shall contain not more than 10% of polyglycerols equal to or higher than heptaglycerol. Not less than 80 and not more than 100 mg KOH/g Not more than 6 mg KOH/g Not more than 3 mg/kg Not more than 5 mg/kg Not more than 1 mg/kg Not more than 1 mg/kg Not more than 10 mg/kg

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2. MATERIALS AND METHODS 2.1. Materials 2.1.1. Enzymes Lipase from Candida rugosa (819 U/mg solid) was purchased from Sigma-Aldrich. Lipases from Rhizopus oryzae (58.4 U/mg solid) and Rhizopus arrhizus (10 U/mg solid) were purchased from Fluka. ―Lipase basic kit‖ was purchased from Fluka. This kit contains lipases from different sources: Aspergillus (culture not specified) (0.2 U/mg solid), Candida antarctica (2.9 U/mg solid), Candida cylindracea (3.85 U/mg solid), Mucor miehei (1.4 U/mg solid), Pseudomonas cepacia (46.2 U/mg solid), Pseudomonas fluorescens (36 U/mg solid), Rhizopus arrhizus (9.18 U/mg solid), Rhizopus niveus (1.7 U/mg solid) and porcine pancreas (20.6 U/mg solid). ―Lipase extension kit‖ was acquired from Fluka. This kit includes lipases from different sources: Aspergillus oryzae (48 U/mg solid), Candida lipolytica (0.0011 U/mg solid), Mucor javanicus (11.6 U/mg solid), Penicillium roqueforti (0.65 U/mg solid), Pseudomonas fluorescens (309 U/mg solid), Rhizomucor miehei recombinant from Aspergillus oryzae (0.51 U/mg solid), wheat germ (0.1 U/mg solid), Chromobacterium viscosum (2711 U/mg solid), Pseudomonas sp. (2324 U/mg solid) and Pseudomonas sp. (Type B) (256 U/mg solid).

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2.1.2. Substrates Ricinoleic acid (80%) was purchased from Sigma-Aldrich. Polyglycerol-3 (PG-3) was kindly donated by Solvay and it is a glycerol oligomer based on an average of three glycerol groups (average MW = 250 g/mol). It contains minimum 80% di-, tri- and tetraglycerol and has very low levels of cyclic by-products. 2.1.3. Immobilisation Reagents and Activators γ-APTES ((3-aminopropyl) triethoxysilane) and glutaraldehyde (25%) were purchased from Sigma-Aldrich. Oleic acid (>58%) was acquired from Riedel-de Haën. Soybean lecithin was of commercial grade from Santiveri S.A., Spain. 2.1.4. Immobilisation Supports Uncoated porous glass beads (PG 75-40, PG 700-400 and PG 1000-400) and acid-washed non-porous glass beads (≤106 μm and 425-600 μm) were acquired from Sigma. Biolita L2.7 and P3.5 (biolite) were a kind gift from Ondeo Degrémont, Bilbao. Chromosorb W (30-60 mesh) and Celite R-643 were from Johns Manville Products. Cationic and anionic exchange resins (Dowex 50×8 and Lewatit MonoPlus MP 64, respectively) were supplied by Fluka.

2.2. Methods 2.2.1. Immobilisation by Covalent Binding The immobilisation process was carried out according to the following steps [15]: Preparation of the carrier: Glass beads were washed in 5% HNO3 at 80-90ºC for 60 min and then rinsed with distilled water and dried in an oven for 24 h at 110ºC.

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Support activation: To 1 g of clean glass beads, 18 ml of water was added along with 2 ml of γ-APTES (10% v/v) and the pH was adjusted to between pH 3 and 4 with 6N HCl. After adjustment, the mixture was placed in a 75ºC water bath for 2 h. The silanized glass was removed from the bath, washed with distilled water and dried overnight in an oven at 110ºC. The resulting product may be stored for later use. Immobilisation on glass-glutaraldehyde: 1 g of silanized glass was made to react in a jacketed column reactor (2.5 i.d. and 30 cm length) with 25 ml of glutaraldehyde 2.5% in 0.05 M phosphate buffer, pH=7. The reactor was equipped with a sinterized glass plate placed 5 cm from the bottom. The solution was recycled for 60 min at room temperature with a peristaltic pump and the glass-glutaraldehyde washed with 25 ml of the same buffer. Enzyme solution (50 ml, 10 mg/ml) was then added to the reactor and the enzyme solution recycled overnight at 4°C. The derivative was then washed three times with 0.1 M phosphate buffer, pH=7. The immobilised derivative was suspended in the same buffer and stored at 4ºC until use. The amount of protein initially offered and in the wash-liquid after immobilisation was determined by Lowry‘s procedure modified by Hartree [16], using bovine serum albumin as standard. The amount of coupled lipase was the difference between the amount of the initial enzyme added and the amount of enzyme in the wash-liquid.

2.2.2. Immobilisation by Physical Adsorption Unless otherwise stated, 1 g of support was mixed with 10 ml of an activator suspension (20 mg/ml) in an erlenmeyer flask and placed in an orbital shaker overnight at room temperature. Three activators were tested: soybean lecithin, ricinoleic acid and oleic acid. One gram of support (as purchased or activated) was washed with 10 ml of distilled water and then transferred to the above mentioned jacketed column reactor. The enzyme solution (10 ml, 10 mg/ml in acetate buffer 0.1 M, pH=5) was then added to the reactor and recycled for one day at 4°C. The immobilised derivative was washed twice with the same buffer and stored at 4°C. When the influence of pH was studied, acetate buffer 0.1 M was used to adjust the pH values to 4, 4.5 and 5 and phosphate buffer 0.1 M was used for pH values of 6 and 7. The amount of immobilised enzyme was determined as described in Section 2.2.1. 2.2.3. Synthesis of Polyricinoleic Acid, PR Polyricinoleic acid was obtained with free and immobilised lipase and several reaction devices were used. 2.2.3.1. With Free Candida rugosa Lipase Preliminary Studies The enzymatic reaction was carried out in a batch reactor (100 ml total volume). Complete mixing was achieved either by orbital shaking or by means of a three-bladed propeller stirrer. The reaction temperature was always kept constant at 40°C [13], which has been found to be the optimal reaction temperature. The heating system was an incubator provided with hot air circulation. The reaction mixture contained 30 g of ricinoleic acid and the appropriate amounts of enzyme and water. Samples were taken and acid value (AV) of the reaction mixture was determined.

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Open Air Reactor Experiments The enzymatic reaction was carried out in an open jacketed batch reactor (100 ml total volume). Complete mixing was achieved by means of a three-bladed propeller stirrer. The reaction temperature was always kept constant at 40ºC [13]. The reaction mixture contained 30 g of ricinoleic acid and the appropriate amount of free Candida rugosa lipase. A certain amount of water is poured over the lipase before the substrate is added to the reactor. The extent of the reaction was followed by means of AV measurements. 2.2.3.2. With Immobilised Candida rugosa Lipase

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Open Air Reactor Experiments The enzymatic reaction with immobilised lipase was similar to that previously described. In this case, the reaction mixture contained 30 g of ricinoleic acid and five grams of immobilised lipase. The only water in the reaction system is that soaked in the support (0.6 ml/g approximately) except in the experiments to study the influence of the initial water content. In all cases, samples were taken from the reactor at given time intervals and AV of the reaction mixture was determined. Vacuum Reactor Experiments For reactions under vacuum, a Parr 5100 series low pressure reactor was used. The reaction vessel (100 ml total volume) is made of glass and is equipped with a circulating jacket to heat the vessel. The reactor head is stainless steel and accommodates the reactor controls and instrumentation. The reactor is equipped with a magnetic drive to provide a trouble free internal stirrer, which is a turbine type impeller. The reactor top also includes a vacuum meter, an internal thermocouple, an internal cooling loop, a rupture disk, a liquid sample valve, a gas inlet valve and a gas release valve. Temperature, stirring speed and positive pressure are managed by a controller. The amount of ricinoleic acid, immobilised lipase and water in the reactor at the beginning of the reaction are the same that those reported for the open air jacketed batch reactor. All the experiments were carried out at 40°C, and the stirring rate was kept constant at a value of 350 rpm. The pressure was set at 160 mm Hg and, in several experiments, a 90 l/h dry nitrogen flow was conducted through the reaction mixture to facilitate water removal. The nitrogen flow was dried by passing through a silica gel column; therefore its relative humidity was zero. 2.2.4. Synthesis of Polyglycerol Polyricinoleate, PGPR Polyglycerol polyricinoleate was obtained with free and immobilised lipase and several reaction devices were used. 2.2.4.1. With Free Lipase Selection Studies The reactions were carried out in a 250 ml jacketed batch reactor at 40°C and the mass transfer was promoted by a powerful four-bladed impeller stirrer, which was used as mixing device. First, the appropriate amount of lipase (see Table 6) was placed to the reactor and 5 ml of distilled water was poured over the lipase. Then, 30 g of PR (AV≤50 mg KOH/g) and 2

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g of PG-3 were added to the reactor, so that the mass ratio PR/PG was 15 (which means that three of the five hydroxyl groups of the polyglycerol could be esterified). As can be seen, the reaction occurs in the absence of solvent and with a limited initial amount of water. Samples were taken from the reactor at given time intervals and the AV of the reaction mixture was determined. All experiments were left to progress for approximately seven days.

Open Air Reactor Experiments The reactions were carried out in the same reactor above described. The reaction mixture was similar to that used for the selection experiments. In this case, 500 mg of Rhizopus arrhizus lipase was dissolved in 5 ml of distilled water and added to the reactor before the substrates. 2.2.4.2. With Immobilised Lipase Open Air Reactor Experiments In this case, 5 g of immobilised derivative was used and the only water in the reaction system was that soaked in the support.

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Vacuum Reactor Experiments The reactor used in this set of experiments was the same utilized for PR synthesis. The amount of ricinoleic acid, immobilised lipase and water in the reactor at the beginning of the reaction were the same as that reported for the open air jacketed batch reactor. The experiments were carried out at 40ºC, and the stirring rate was kept constant at 450 rpm. The pressure was set at 160 mm Hg and 90 l/h dry nitrogen was passed through the reaction mixture to facilitate water removal (the nitrogen flow was dried by passing through a silica gel column, so that its relative humidity was zero). 2.2.5. Measurement of the Reaction Extension The acid value (AV) [17], which represents the number of milligrams of potassium hydroxide necessary to neutralize free acids in 1 g of sample, was used as an index to show the reaction extension. Here, the AV corresponds to the free carboxyl group concentration in the reaction mixture, which decreases due to the autocondensation of the ricinoleic acid and/or the esterification of polyricinoleic acid (AV≤50 mg KOH/g) with polyglycerol-3. 2.2.6. Measurement of the Water Content Water content was measured in the reactor samples with a Karl-Fischer automatic titrator (701 KF, Metrohm), using Hydranal® composite 5, from Riedel-De-Häen. 2.2.7. Recovery of the Immobilised Derivative When immobilised derivatives were tested for reusability, the reactor content was placed in a sinterized glass filter (Pirex®, number 0) to separate the derivative from the product. After 8 hours at room temperature the immobilised derivative was placed in the reactor for a new reaction cycle.

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3. RESULTS AND DISCUSSION 3.1. Condensation of Ricinoleic Acid to Obtain Polyricinoleic Acid by Candida rugosa Lipase

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After a conscientious bibliographical search, Candida rugosa lipase has been selected to catalyse the autocondensation reaction of ricinoleic acid to obtain the estolide, also called polyricinoleic acid, PR [13].

3.1.1. Preliminary Studies with Free Candida rugosa Lipase Because the synthesis of ricinoleic acid estolide is a dehydration–condensation reaction, the reaction reaches equilibrium and stops when the water content in the reaction mixture increases as a result of the water formed during the reaction. It has been reported [13] that the polarity of the bulk reaction mixture becomes lower as condensation progresses and most water molecules associated to the polar groups of ricinoleic acid are now free to maintain the configuration of lipase. Therefore, the optimal water content decreases during the reaction course and water must be removed from the reaction mixture. By comparing experiments conducted at similar initial conditions of enzyme concentration, water content and stirring speed, but using different dehydration methods (spontaneous evaporation, a vacuum aspirator and hot air current), hot air was found to be the best way of removing excess water and obtaining estolides with a high degree of condensation (data not shown). First, the influence of a hot air current on the water content of ricinoleic acid was studied. For this, 30 g of ricinoleic acid were mixed in open flasks with different volumes of water to give initial water concentrations between 10,000 and 510,000 ppm. Then, they were placed in an incubator with an orbital shaking apparatus. The incubator was thermostated to 40ºC using hot air circulation. The total water content of the mixture was followed by taking samples and titrating using the Karl–Fisher method (Section 2.2.6). The results show that, in all cases, most of the water was eliminated during the first 2–10 h, depending on the initial water concentration. After 24 h, the water content had stabilized at about 2500 ppm in all the experiments. The presence of a small amount of water is necessary for the expression of the enzyme activity because water is essential for maintaining the configuration of the enzyme in the proper form [18]. On the other hand, too high a water content might displace the equilibrium towards the hydrolysis reaction. Therefore, the influence of the initial water content on the condensation reaction should be studied. A set of experiments was carried out varying the initial concentration of water in the reaction mixture between 4000 and 510,000 ppm. The two lowest water contents were achieved by drying commercial ricinoleic acid (water content 10,000 ppm). All the reactions were carried out with 30 g of ricinoleic acid and 200 mg of lipase from Candida rugosa. The appropriate amounts of water were added when initial water contents higher than 10,000 ppm were assayed. It can be observed from Figure 2 that the ricinoleic acid condensed even when the minimum initial water content (4000 ppm) was used. This result is in accordance with the only previous report found in the literature, which mentions that the minimum water content required for expressing lipase activity lies between 1800 and 4200 ppm for ricinoleic acid estolide biosynthesis [13].

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Figure 2. Kinetic of the condensation reaction at different initial water concentrations. Reaction conditions: enzyme concentration, 6.67 mg E/g ricinoleic acid; initial water content, 4000 ppm (♦), 5800 ppm (◊), 10,000 ppm (▲), 77,000 ppm (Δ), 110,000 ppm (■), 310,000 ppm (□) and 510,000 ppm (●).

As the water content was increased from 4000 to 77,000 ppm, so the decrease in AV became more pronounced. This was because the initial water concentration in the reaction mixture was not sufficient to maintain the enzyme in a catalytically active conformation and the enzymatic activity increased with increasing concentrations of water. On the other hand, when the water concentrations were varied between 77,000 and 310,000 ppm, no significant changes in the AV/time slopes were observed. In these four experiments, the water in the reaction mixture enabled the enzyme to catalyse the synthetic reaction at a very high reaction rate. However, when 15 ml of water were added to the reactor (to provide a water concentration of 510,000 ppm) the reaction rate decreased. In this case, the high water concentrations favoured the reverse reaction (law of mass action). From these experiments it was not possible to establish the optimum water content because high reaction rates were observed for a wide range. However, it can be assumed that the addition of water to the commercial ricinoleic acid benefits the condensation reaction as long as the initial amount of added water is lower than 0.5 ml H2O/g ricinoleic acid (initial water concentration lower than 510,000 ppm). These results agree with previously published findings [13] only from a qualitative point of view because these authors reported optimum water contents noticeably lower than ours (between 3000 and 17,000 ppm). This difference could be explained by differences in the ricinoleic acid used as substrate of the enzymatic reaction. Another reason could be the fact that the above-mentioned authors made use of an immobilised lipase in a ceramic support and, in this microenvironment, the amount of water required by the enzyme to express its catalytic activity might be lower. In view of those results, a water content of 77,000 ppm is considered adequate for an enzyme concentration of 6.67 mg E/g ricin. Later another set of experiments was conducted to study the influence of the enzyme concentration on the reaction rate. Due to the main role of water is to maintain the enzyme configuration, it may be thought that an increasing amount of enzyme should accompany an increasing amount of water. In all cases the water concentration was maintained in the range where the reaction rate was maximal (see Figure 2). Therefore, another series of experiments

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(B)

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Figure 3. Time course of the condensation reaction. Conditions: () [enzyme] = 6.67 mg E/g ricin; [H2O] = 77,000 ppm; (■) [enzyme] = 13.33 mg E/g ricin; [H2O] = 144,000 ppm; (▲) [enzyme] = 20.00 mg E/g ricin; [H2O] =211,000 ppm. (A) Orbital shaker and (B) three-bladed propeller stirrer.

was planned, in which both enzyme and water concentrations were simultaneously increased. The results of this experimental series are shown in Figure 3A. As can be seen, a substantial improvement in the reaction rate was achieved when added water and enzyme concentrations were doubled but no additional enhancement was observed when the concentration values were increased threefold. As a consequence, it was concluded that no faster processes could be expected by increasing the biocatalyst concentration. Even at the 40ºC used, the viscosity of the reaction medium was quite high, and it may be thought that the reaction process was controlled by mass transport phenomena. Another experimental series was carried out with the same water and lipase concentrations as in the previous series but using a three-bladed propeller stirrer instead of orbital shaking for mixing. The results are shown in Figure 3B, where it can be seen that the reaction rate improves only in the case of the lowest enzyme and water concentrations. Comparing both figures, it can be concluded that when the lower enzyme concentration was used the condensation reaction progressed under diffusional control because the reaction rate was improved by more efficient mixing. However, when larger enzyme concentrations (double and triple) were assayed no improvement in reaction rate was observed by changing the mixing device due to kinetic control. Therefore, the optimum enzyme concentration lies between 6.67 and 13.33 mg E/g

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ricin. These results could not be compared with others because no previous studies have been found in the literature. It has been reported that the acid value of ricinoleic acid estolide prepared by conventional chemical methods is 40 mg KOH/g [2] and that this AV is only attainable by using immobilised lipase and never with free enzyme [19]. The acid value is related with the degree of polymerisation and each particular application of the estolide will require a different AV value. Using the reactor configuration described and the best reaction conditions determined (water and enzyme concentrations), an AV value of 65 mg KOH/g was reached after 48 h. This reaction time is shorter than that reported previously [19] where ricinoleic acid estolides with AV of 60 mg KOH/g were produced with free lipase after 150 h of reaction. These results can probably be improved by using the lipase in an immobilised form.

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3.1.2. Immobilisation of Candida rugosa Lipase In order to improve the results obtained and due to the well-known advantages of the immobilised enzymes, the following studies were devoted to obtain an immobilised derivative of Candida rugosa lipase with high activity and stability which could be used as an efficient catalyst of the reaction under study. 3.1.2.1. Choice of the Immobilised Derivative It has been described that adsorbed lipase on a ceramic carrier SM-10 [13] and the commercial immobilised lipase Novozym® 435 are suitable for producing ricinoleic acid estolide. However, the difficulty of acquiring the former support and the small size and low density of the latter, which hinders its separation from the reaction mixture, led the authors to test different immobilisation matrices in an attempt to obtain an immobilised derivative, which could be successfully used to catalyse the production of ricinoleic acid estolide. Eight inorganic supports (two types of biolite, Celite R-643, Chromosorb W, non-porous glass beads of two particle sizes and porous glass beads of different pore sizes) and two organic carriers (cationic and anionic exchange resins, Dowex 50×8 and Lewatit MonoPlus MP 64, respectively) were used. Twelve different immobilised derivatives were obtained, six of them by physical adsorption and the other six by covalent coupling via the amino groups of the enzyme. Immobilisation on glutaraldehyde-activated aminopropyl glass beads was selected because it has been widely used by the authors with different enzymes [14, 20] and has been shown to be very versatile. The results obtained are shown in Table 2 where percentages of immobilised protein and protein contents are summarized. It is important to note that these values are based on the protein content provided by Lowry‘s method [16], which showed that the commercial lipase contained only 15% protein. The best results were obtained when porous glass was used as immobilisation matrix and covalent binding as coupling method. In these cases enzyme loading increased as the pore size became smaller because of the greater internal surface available for immobilisation. The percentage of immobilised lipase obtained by physical adsorption on Lewatit MonoPlus MP 64 was higher than those obtained with porous glass because five times less enzyme was offered for immobilisation, so that enzyme loading (mg E/g support) was noticeably lower. Celite R-643 was also shown to be suitable for Candida rugosa lipase immobilisation.

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Table 2. Coupling parameters for the immobilisation of Candida rugosa lipase in different supports

Support

Immobilisation method

Immobilised protein (%)

Biolita L2.7

Adsorption Covalent binding

– 0.64 – 27.60 –

Enzyme loading (mg E/g support) – 0.48 – 4.14 –

0.89

0.67

–

–

41.73 28.65 22.83

31.30 21.49 17.12

–

–

47.20

7.08

Biolita P3.5 Celite R-643 Chromosorb W Non-porous glass 425–600 µm Non-porous glass 91–107 µm PG 75-400 PG 700-400 PG 1000-400 Cationic exchange resin Dowex 50×8 Anionic exchange resin Lewatit MonoPlus MP 64

Adsorption

Covalent binding

Adsorption

200

AV (mg KOH/g)

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The five above mentioned immobilised derivatives were used to catalyse the polymerization reaction of ricinoleic acid following the procedure described in Section 2.2.3.2. The results obtained are shown in Figure 4.

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Time (h) Figure 4. Change in acid-value as a function of time for estolide synthesis catalysed by five different immobilised derivatives. ( ▲ ) Celite R-643, ( ■ ) PG 75-400, ( x ) PG 700-400, ( ● ) PG 1000-400, (  ) Lewatit MonoPlus MP 64.

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It can be seen that there was a large difference between the activity of the derivative obtained on the anion exchange resin and the activity of other derivatives. In the case of the anion exchange resin, the acid value dropped from 180 to 50 in 150 h while the best result of the other derivatives was a fall to 136 in 285 h (porous glass 75–400). Therefore, the immobilised lipase obtained by physical adsorption onto Lewatit MonoPlus MP 64 was chosen for further studies.

3.1.2.2. Choice of Support Activator It has been described [21] that esterifying activity of lipase is higher when the enzyme is immobilised on a carrier previously activated with phospholipids, fatty acids or fatty acid esters. In this case, lipase is adsorbed at the water–fat interface on the surface of the support and the enzyme activity can be efficiently utilized. Table 3 shows the results obtained in the immobilisation processes with and without activators, in terms of percentage of immobilised protein and enzyme loading. Similar results were obtained for all the immobilised derivatives, pointing to the null effect of activator on the immobilisation process as described elsewhere [21]. Table 3. Coupling parameters for the immobilisation of Candida rugosa lipase by physical adsorption on Lewatit MonoPlus MP 64 using different support activators Activator

Enzyme loading (mg E/g support) 7.08 7.93 5.34 7.32

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None Soybean lecithin Ricinoleic acid Oleic acid

Immobilised protein (%) 47.20 52.86 35.60 48.80

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Time (h) Figure 5. Influence on the evolution of acid value with time of the addition of three different activators during the lipase immobilisation process. ( ■ ) None, ( ▲ ) ricinoleic acid, (  ) oleic acid, ( x ) soybean lecithin.

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The four immobilised derivatives were tested for activity in a batch reactor, following the procedure described in Section 2.2.3.2. The results obtained are shown in Figure 5, where the beneficial effect on the enzymatic activity provoked by ricinoleic acid and soybean lecithin is pointed out. Estolide with an AV close to 40 mg KOH/g was obtained after 100 h of reaction. Slightly better results were achieved when soybean lecithin was used, and so this phospholipid was chosen as activator for further experiments. It is important to emphasize that soybean lecithin is also the activator used previously by Y. Yoshida et al. in the only biocatalytic process described in the literature to obtain ricinoleic acid estolide [13]. However, as mentioned before, there are no papers on the use of other activators, so we can just suppose that the authors used their intuition or that they have previous unpublished studies.

3.1.2.3. Influence of Enzyme Concentration and pH The support Lewatit MonoPlus MP 64 is a weakly basic anion exchange resin and its nature suggests that any adsorption of proteins would be governed by electrostatic forces. Therefore, changing the pH value should have a large impact on adsorption. On the other hand, the carrier is based on a styrene-divinylbenzene copolymer which together with the phospholipids used as activator, could confer a certain hydrophobic character to the support. In this case, the adsorption could be controlled by hydrophobic interactions and the amount of protein adsorbed would not be significantly influenced by pH changes in the protein solutions used for adsorption. In order to explain which interaction predominates in the adsorption of lipase on Lewatit MonoPlus MP 64, several immobilised derivatives were obtained using lipase solution concentrations varying between 2 and 30 mg E/ml (which corresponds to 0.3–4.5 mg Lowry‘s protein/ml) dissolved in different buffers in a pH range from 4 to 7. The results obtained are shown in Figure 6. It can be seen that for each pH value assayed, an increase in enzyme concentration up to 4 mg E/ml enhanced the driving force for the adsorption and increased the amount of protein adsorbed correspondingly. At pH≥6 and when protein concentration was higher than 4 mg E/ml, the increase slowed down gradually. Lipase adsorption was improved at pH≤5 and, in this case, the amount of enzyme adsorbed continuously increased as protein concentration increased (up to 30 mg E/ml), which means that higher enzyme concentrations could be used, resulting in an immobilised derivative of 14 mg E/g support. In the article of Yoshida [13] describing estolide production, two different derivatives with 60 and 120 mg E/g support were obtained, the authors considering that the entire enzyme is attached to the support during the immobilisation process. The maximum capacity of the polypropylene carrier used by Gitlesen et al. [22] was about 220 mg E/g support; however, this parameter could not be verified because we found difficulties when trying to dissolve higher enzyme concentrations in the immobilisation buffer. Nonetheless, the maximum capacity of the Lewatit MonoPlus MP 64 support at pH≤5 was not reached. Other authors obtained immobilised derivatives of lipase with a lower (0.17 mg E/g [23]; 0.071–0.129 mg E/g [24]; 0.25–6.5 mg E/g [25]) or higher (150–1000 mg E/g [26]) enzyme contents.

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Enzyme loading (mgE/g support)

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Figure 6. Influence of buffer pH and the offered protein concentration on the enzyme loading for the immobilisation of lipase on Lewatit MonoPlus MP 64. (  ) pH=4, ( ■ ) pH=4.5, ( ▲ ) pH=5, ( x ) pH=6, (  ) pH=7.

In order to compare the specific activity of the immobilised lipase and the free enzyme, two experiments were carried out using the same amount of protein as catalyst; that is, one experiment was performed with the immobilised derivative and the other was prepared with the same amount of protein adsorbed on the support. In Figure 8 the evolution of the acid values measured in both reactors are represented. Lipase immobilisation is clearly affected by pH changes in the absorption solution, suggesting that electrostatic interactions are the main driving force in the adsorption of lipase by Lewatit MonoPlus MP 64. The isoelectric point (pI) of Candida rugosa lipase is 4.6–4.7 [27], so when the pH is lower than pI, the enzyme takes on a positive charge. Since the absorption process is favoured at pH values lower than 5.0, the carrier has a negative charge, revealing that exchange sites of the support have a pK of less than 4.0. For pH values between 4.6 and 5.0, both enzyme and carrier have negative charges and in such a situation the lipase could possess localized areas of high positive charge on its surface (i.e. discrete binding sites) or other forces (hydrophobic forces) could play a role in the binding process. On the other hand, the shape of the adsorption isotherms shows a sigmoidal shape at pH 7, indicating that enzyme–enzyme forces are more important than the support–enzyme forces, and the typical Langmuir trend at pH≤5 indicates that at the carrier surface the support– protein interactions are stronger than protein–protein ones [28]. No trend is clear at pH 6 (data not shown).

3.1.2.4. Effect of Mixing in the Immobilisation Process It has been described that lipase adsorption to solid surfaces follows several steps. The lipase molecule in the bulk phase must be transported to the surface, either by convection or diffusion. Even in well-stirred systems there exists a stagnant layer close to the surface that must be penetrated by diffusion. The lipase is then adsorbed at the solid surface at a given

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Time (h) - Column reactor 5 10 15 20

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Figure 7. Time course of protein absorption in the cool room (bottom time scale) and in the column reactor (top time scale). ( ● ) Cool room, ( ○ ) column reactor.

rate. After adsorption, macroscopic or microscopic rearrangements in protein structure can occur. Desorption of adsorbed protein is not common and the process is apparently irreversible [22]. In a standard experiment of protein adsorption, the activated support is placed in a jacketed column reactor and enzyme solution is added and recycled for 48 h at 4ºC. Although adsorption processes are usually slow, 2 days seemed to be sufficient to reach equilibrium. To check whether the adsorption process had been completed and to optimize the immobilisation time, an experiment was carried out in which samples were taken from the supernatant and analysed for enzyme. In parallel, the same amounts of activated support and enzyme solution were poured together into an erlenmeyer flask and the mixture was placed in the cool room at 4ºC. This supernatant was also sampled and analysed for protein. The results of both experiments are shown in Figure 7, in which enzyme concentration in the supernatant is represented against time (note that time scales are different). Although the final enzyme concentration was exactly the same in the column reactor and erlenmeyer flask, the equilibrium concentration in the former device was reached after less than 10 h, while 2 days were necessary to complete the immobilisation step in the cool room.

3.1.2.5. Comparison of Free and Immobilised Lipase To recapitulate, the immobilisation procedure of Candida rugosa lipase was optimized and the best results, taking into consideration enzyme consumption and the activity of the resulting derivative, were obtained when 50 ml of enzyme solution (10 mg/ml in acetate buffer 0.1 M, pH 5.0) was mixed with 5 g of the anion exchange resin Lewatit MonoPlus MP 64 previously activated with soybean lecithin, and recycled in a column reactor for 10 h at 4ºC or left to stand in the cool room for 2 days. Under these conditions an immobilised derivative with almost 8 mg E/g support was obtained. Furthermore 5 g of resin soaks up 3 ml of water.

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AV (mg KOH/g)

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Figure 8. Change in acid value as a function of time for estolide synthesis catalysed by free and immobilised lipase. (  ) 40 mg immobilised protein, ( ■ ) 40 mg free protein.

After 24 h, the AV of the reaction with native lipase decrease to half of its initial value, while for the reaction catalysed with the immobilised enzyme the AV decrease to one third of its initial value. In addition, the final AV attained (close to 40 mg KOH/g) was noticeably lower when immobilised lipase was used as catalyst. In the work of Yoshida et al. [13], the estolide produced per unit weight of lipase was 425 g estolide/g free enzyme, when immobilised derivative (60 mg E/g) was used in five consecutive batches. In our case, 750 g of estolide are obtained per gram of free enzyme in an only run, and this amount may be increased if the immobilised derivative is evaluated and proved for reuse. A similar comparison could not be established with the results of Kelly and Hayes [29] because they use a commercial immobilised lipase and the derivative enzyme content is not revealed.

3.1.2.6. Reuse of the Immobilised Derivative Immobilisation provides an attractive opportunity for the multiple use of the same enzyme. In order to establish the reusability of our immobilised derivative, successive polymerization reactions were planned. The purpose of this experiment was to assess the multiple use of the immobilised lipase by examining the evolution of AV with time in several consecutive experiments. The results are shown in Figure 9. During the first 24 h, AV decreases almost 120 units in the first use, 70 units in the second run, and only 48 in the third run. However, after 7 days, the reached AV in the second use of the derivative is almost the same than that in the first utilization. In order to ascertain whether this activity loss is due to enzyme desorption or inactivation an additional experiment was carried out. After 24 h in a normal polymerization process with immobilised derivative, the reactor content was divided in two halves so that the support remains in the original reactor. Then 15 g of ricinoleic acid was added to both reactors and the evolution of AV was registered. In the original reactor a decrease of AV with time was observed and in the second reactor no variation of AV with time was detected (data not shown). In consequence it can be affirmed that activity loss in reuse experiments was caused by enzyme deactivation.

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AV (mg KOH/g)

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50

100 Time (h)

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Figure 9. Synthesis of estolide by repeated batch operation. (  ) First run, ( ■ ) second run, ( ▲ ) third run.

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Table 4. Operating conditions for the lipase-catalysed synthesis of ricinoleic acid estolides in the open air jacketed batch reactor Run 1 2 3 4 5 6 7 8 9

IME (g) 2.5 5 10 5 5 5 5 5 5

Water Soaked Soaked Soaked Soaked Soaked Soaked dry +0.5 ml +5 ml

Temperature (◦C) 40 40 40 50 45 40 40 40 40

3.1.3. Production of Polyricinoleic Acid with Immobilised Candida rugosa Lipase Once we have obtained an immobilised derivative with the appropriate characteristics to be used in the synthesis of ricinoleic acid estolides, several experiments were performed in the open air jacketed batch reactor in order to study the influence of three operating variables on reaction. The experimental conditions in which the reactions were carried out are summarized in Table 4. 3.1.3.1. Influence of the Amount of Enzyme For the synthesis of polyricinoleic acid, the influence of different amounts of added immobilised lipase on the degree of polymerization of the estolides was studied. As shown in Figure 10, when the amount of immobilised enzyme (IME) added to the reactor was

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increased, the reaction progressed faster and a lower acid value was reached. The differences between the acid values reached led us to think that these were not equilibrium values but that the reaction stops because the enzyme is not active anymore. However, changes in reaction rate and final acid value were more noticeable when the enzyme amount varied from 2.5 to 5 g than when the enzyme amount was increased to 10 g, meaning that in the last case more enzyme than necessary was being used. On the other hand, 10 g of IME in 30 g of ricinoleic acid was a ratio too high to be easily handled. In consequence 5 g of immobilised derivative was considered as appropriate for further experiments.

3.1.3.2. Influence of the Temperature The effect of the temperature on the reaction course was investigated. Temperature was seen to influence the enzymatic reaction rate, the enzyme stability, the velocity of water evaporation from the reaction medium and its viscosity. The lowest temperature chosen was fixed at 40ºC, which is the optimum temperature of Candida rugosa lipase [13]. Below this temperature stirring the reaction mixture became difficult because of the high viscosity of the ricinoleic acid and its estolides. The upper temperature limit was fixed at 50ºC to avoid enzyme denaturalization. When the temperature was raised from 40 to 50ºC, the viscosity values of ricinoleic acid and polyricinoleic acid fell by 35% (data not shown). Figure 11 shows the evolution with time of the acid value at each temperature. As can be seen, this variable had little influence on the reaction course within the range studied, probably because it was not broad enough and because the different effects (viscosity, reaction rate and enzyme deactivation) are compensated. Nevertheless, a slightly unfavourable effect could be observed at high temperature. Therefore, for further experiments 40ºC was chosen as optimum temperature.

AV (mg KOH/g)

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Time (h) Figure 10. Influence of the amount of enzyme on the evolution of acid value with time for the PR synthesis in the open reactor. (  ) 2.5 g IME, ( ■ ) 5 g IME, ( ▲ ) 10 g IME.

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AV (mg KOH/g)

200 150 100 50 0 0

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100 Time (h)

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Figure 11. Change in acid value as a function of time for estolide synthesis performed at different temperatures in the open reactor. (  ) 40ºC, ( ▲ ) 45ºC, ( ■ ) 50ºC.

3.1.3.3. Influence of the Initial Water Content As described above, water plays multiple roles on lipase catalysed esterifications performed in non-conventional media. It is widely known that water is absolutely necessary for the catalytic function of enzymes because it participates, directly or indirectly, in all noncovalent interactions that maintain the conformation of the catalytic site of enzymes [10–12]. However, it has been found that the amount of water necessary for enzyme activity might be very small and, in the case of lipase, just a few layers around the enzyme surface are needed [30]. In addition, the water content affects the equilibrium in esterification/hydrolysis reactions as well as the distribution of products in the media [31]. Particularly, in case of esterifications, lower conversions are achieved as the water content increases. In the light of the above considerations, a study on the optimal initial amount of water in the reactor was deemed necessary. With this purpose, four experiments were carried out using the immobilised derivative as obtained (soaked), adding different amounts of water, and drying the derivative under vacuum at room temperature before use. The time course of these experiments is shown in Figure 12 where the acid value is represented against operation time. With these experiments it was demonstrated that an optimum in the initial water content exists, although this optimum seems to be quite wide. The same results were obtained when derivative was used as obtained and when small amounts of water were added. However, drying the derivative or adding higher amounts of water led to a lower initial rate (specially the high water content) and a higher final acid value.

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AV (mg KOH/g)

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Time (h)

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Figure 12. Influence of the initial water content on the evolution of acid value with time for the PR synthesis in the open reactor. ( ▲ ) Wet resin, ( ● ) dry resin, ( ■ ) addition of 0.5 ml of water, (  ) addition of 5 ml of water.

All the experiments described above were carried out simultaneously, in an air open tank reactor, within a month of each other. However, when the results were tested for reproducibility, great discrepancy between them was observed, as shown in Figure 13, where a variation of 30% in the AV value was obtained for experiments carried out in different seasons. Measurements of the water content of the reaction medium revealed that, at 40ºC, water continuously evaporated and that after 48 h (approximately) the reactor water content was independent of the initial water content and mainly dependent on environmental relative humidity. The air conditioner/heat pump equipment installed in the laboratory stabilizes the relative humidity at 70% in summer (air conditioner) and at 20% in winter (heat pump). These values correspond with equilibrium water contents of 3600 and 1000 ppm, respectively, which are the cause of the discrepancies in the results obtained in the open air reactor. Obviously, this poor reproducibility of the results is unacceptable if the process is to be applied on an industrial scale. Therefore, the remaining experiments were carried out in the vacuum reactor described in Section 2.2.3.2 in which the synthesis of ricinoleic acid estolides can be conducted in a closed system with controlled atmosphere, a suitable level of stirring and low pressure.

3.1.3.3. Influence of Pressure and Water Content First, the influence of pressure on PR synthesis was analysed. The results obtained in the experiment carried out at a constant pressure of 160 mm Hg are illustrated in Figure 14, where it can be observed that, after 50 h of reaction, the acid value of the estolide obtained under vacuum was only 70 mg KOH/g, which is noticeably higher than the best result obtained in the open air reactor. It was thought that water removal could be improved by using a vacuum [13], but at the end of the experiment (70 h), it was still higher than 30,000 ppm. This might be attributable to the partial condensation of the evaporated water (which consequently drops into the reaction mixture) due to the geometry of the reactor top plate.

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AV (mg KOH/g)

200

150

100

50

0 0

50

100 Time (h)

150

Figure 13. Comparison of two experiments of PR synthesis carried out in different environmental relative humidity conditions. (  ) 20% relative humidity, ( ■ ) 70% relative humidity.

200 175

AV (mg KOH/g)

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In order to enhance dehydration, a current of dry nitrogen was passed through the reaction mixture and the results obtained in that experiment are also plotted in Figure 14. As can be seen, a slight improvement in the degree of condensation was achieved, compared with the synthesis carried out without dry air, although the acid value reached at the end of the experiment was still too far from that obtained in the open air reactor. In this case, the reactor water content was only 1500 ppm after 24 h and the reaction medium was almost anhydrous (0 ppm Karl-Fisher) at the end of the process. These results demonstrate the high rate of dehydration that occurred and consequently the reaction stopped probably because the enzyme had insufficient water to maintain the active form and therefore, estolides with very high acid value were synthesized.

150 125 100 75 50 25 0 0

50

Time (h)

100

150

Figure 14. Influence of vacuum and dehydration with dry air on the evolution of acid value with time for the PR synthesis. ( ■ ) open air reactor, ( ● ) 160 mm Hg, ( ▲ ) 160 mm Hg with dry nitrogen.

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AV after 48 hours (mg KOH/g)

100 80

no air

21 h 2.5 h

60

4h

5h

7h

40 20 0 1

Drying time (h)

Figure 15. Acid values obtained after 48 h of PR synthesis in the vacuum reactor, at 160 mm Hg, for different values of drying time.

Table 5. Summary of results obtained in the experiments carried out to optimise drying time. The operation time has been 48 hours in all the assays

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Drying time (h) No air 3 4 5 7 9 21

(AV)/h (mg KOH/g h) 1.81 1.94 1.96 2.08 2.21 2.10 1.93

[H2O]/h (ppm/h) 403 1567 1689 1747 1787 1824 1912

To study the influence of dry gas intake, several experiments were performed in the vacuum reactor passing the dry gas for different lengths of time, ranging from 2.5 to 21 h. Samples were taken from the reactor at given time intervals and both the acid value and water content of the reaction mixture were determined. The results obtained are shown in Figure 15, where the acid values reached at 48 h are plotted in a bar chart. It can be observed that the highest degrees of esterification were reached when drying times of 5, 7 and 9 h were assayed. With the objective of analysing these results in detail, the ―average reaction rate‖ (expressed as the acid value variation per hour of reaction) and the ―average rate of water removal‖ were calculated and are summarized in Table 5. As was expected, higher drying times led to higher rates of water removal; however, long drying periods promoted total water removal from the reaction mixture, which supposedly provoked a slight dehydration of lipase and the consequent loss of activity. This explains the

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lower reaction rate calculated when drying times of 21 and 9 h were assayed in comparison with that obtained for the experiment carried out with a drying time of 7 h. These results agree with the revised literature, in which some authors have reported difficulty in removing water from the reaction medium without dehydrating the enzyme, in an attempt to obtain high conversion yields in esterification reactions catalysed by immobilised lipase [10], as soon as the importance of maintaining a minimum water content for the expression of enzymatic activity [13]. In relation with the reaction rate, the best results were obtained for a drying time of 7 h, which is sufficient to guarantee a good rate of water removal, without provoking the elimination of all the water contained in the reaction mixture. In this case, the final content of water in the reactor was 3000 ppm, which is suitable to maintain the lipase perfectly hydrated and, consequently, no loss of activity was observed [13]. For the other drying times assayed, lower rates of water removal were obtained, and the final content of water in the reactor was higher. Hence, esterification reaction progressed more slowly, and a higher acid value was reached. The ricinoleic acid estolides obtained under the above-described reaction conditions had a similar degree of condensation to those synthesized in an open air reactor when the environmental relative humidity was 20%, with the advantage that, in this case, results are totally reproducible.

3.2. Esterification of Polyricinoleic Acid with Polyglycerol to Obtain PGPR

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Once we are able to produce polyricinoleic acid with the appropriate acid value (AV≤50 mg KOH/g) it is used as substrate of the second reaction, i.e., the esterification of PR with polyglycerol.

3.2.1. Screening and Selection of Lipases for the Enzymatic Production of PGPR As described above, lipase from Candida rugosa was used to carry out the autocondensation of ricinoleic acid to obtain the estolide, which is the first step in PGPR synthesis. Obviously, it would be very convenient if the same lipase could serve as catalyst for the two reaction steps. Therefore, the first lipase used to catalyse the esterification reaction between the polyricinoleic acid and polyglycerol was the lipase from Candida rugosa. The obtained results were not satisfying; after 48 hours the decrease of acid value was only 12 mg KOH/g (AV≈42 mg KOH/g at the beginning of the reaction), and the final acid value reached was 30 mg KOH/g. This value is very far from specific purity criteria for PGPR established by the European Commission Directive 98/86/EC [14], in which it is reported that for PGPR to be used as food additive, the acid value must not be higher than 6 mg KOH/g. 3.2.1.1. Selection of Lipases In light of this result, the lipase from Candida rugosa was considered unsuitable for PGPR synthesis and therefore others lipases were assayed for this purpose.

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Enzyme

Source

B A S I C

1 2 3 4 5

K I T

6 7 8 9

Aspergillus sp. Candida antarctica Candida cylindracea Mucor miehei Pseudomonas cepacia Pseudomonas fluorescens Rhizopus arrhizus Rhizopus niveus Porcine pancreas

10

Aspergillus oryzae

E X T E N S I O N

11 12 13 14 15

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16 K I T

17 18 19 20

Candida lipolytica Mucor javanicus Penicillium roqueforti Pseudomonas fluorescens Rhizomucor miehei recombinant from Aspergillus oryzae Wheat germ Chromobacterium viscosum Pseudomonas sp. Pseudomonas sp. (Type B) Rhizopus oryzae

Activity (U/mg solid) (as declared by the manufacturer) 0.201 2.92 3.852 1.42 46.22 362

Added amount (mg) 100 50 1000 100 100 50

3

9.18 1.74 20.65

500 1000 1000

482

100

0.0011 5

2

1000

11.6 0.655

500 500

3092

50

0.512

50

0.11

500

27112

25

23242

10

2566

50

58.4

7

500

1 Unit corresponds to the amount of enzyme which liberates 1 mol acetic acid per minute at pH 7.4 and 40ºC, using triacetine as substrate. 2 1 Unit corresponds to the amount of enzyme which liberates 1 mol oleic acid per minute at pH 8.0 and 40ºC, using triolein as substrate. 3 1 Unit corresponds to the amount of enzyme which liberates 1 mol butyric acid per minute at pH 8.0 and 40ºC, using tributyrin as substrate. 4 1 Unit corresponds to the amount of enzyme which liberates 1 mol fatty acid from a triglyceride per minute at pH 7.7 and 37ºC, using olive oil as substrate. 5 As (4) but at pH 8.0. 6 1 Unit corresponds to the amount of enzyme which liberates 1 mol oleic acid per minute at pH 8.0 and 37ºC, using cholesteryl oleat as substrate. 7 As (4) but at pH 7.2. 1

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Figure 16. Evolution of acid value with time for the PGPR synthesis catalysed by free lipases from different sources. (A) ♦ Aspergillus sp.; ■ Candida antarctica; × Candida cylindracea; ▲ Mucor miehei; + Pseudomonas cepacia. (B) ♦ Candida lipolytica; ■ Mucor javanicus; × Penicillium roquefortii; ▲ Pseudomonas fluorescens (300 U/mg solid); + Rhizomucor miehei. (C) ♦ Pseudomonas fluorescens (40 U/mg solid); ■ Porcine pancreas; × Rhizopus niveus; ▲ Rhizopus arrhizus; + Aspergillus oryzae. (D) ♦ Wheat germ; ■ Chromobacterium viscosum; × Pseudomonas sp. (1200 U/mg solid); ▲ Pseudomonas sp Type B (≥160 U/mg solid); + Rhizopus oryzae.

A further twenty lipases from different sources were used and the corresponding experiments of PGPR synthesis were performed as described in Section 2.2.4.1. Table 6 shows the lipases tested, their specific activities (as declared by the manufacturer) and the amounts of protein used in each experiment. It is important to note that many of the lipases were part of two kits and the amount available was limited. In such cases, the total available protein was added to the reactor. The evolution of the acid value with time for the enzymatic production of PGPR with the above mentioned lipases is plotted in Figure 16 (A to D). In a first selection, eight lipases were rejected because they were not able to reach acid values lower than 15 mg KOH/g in seven days; they are lipases from Aspergillus sp., Candida antarctica, Candida cylindracea, Candida lipolytica, Penicillium roqueforti, porcine pancreas, Rhizopus niveus and wheat germ. The lipase from wheat germ exhibited a particular behaviour. When it was tested to produce PGPR, the acid value of the reaction mixture increased, which indicates that polyricinoleic acid is being hydrolysed and, therefore, under the experimental conditions, the hydrolytic activity of this lipase is greater than its synthetic activity. None of the twelve remaining lipases were able to produce a PGPR with an acid value lower that 6 mg KOH/g, which is the requirement of the European Commission Directive

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[14], although we considered that, after applying appropriate optimization procedures, one or more of these enzymes might be able to efficiently catalyse the enzymatic synthesis of PGPR. The twelve chosen lipases were all from microbial sources, being some 1,3-specific and others ―random‖ lipases. It was thought that any acid value decay in the reaction mixture might be due to two possible reactions: (i) the synthesis of estolides with a higher polymerization degree and (ii) the esterification of polyricinoleic acid with polyglycerol-3 (the desired process). It has been described in the literature that the enzymatic synthesis of estolides can only be successfully catalysed by lipases that lack 1,3-positional selectivity [18, 23, 32], so that lipases from Chromobacterium viscosum and from Pseudomonas (which are ―random‖ lipases and show the best results, Figure 16 D) should be capable, in theory at least, of catalysing the first step of the enzymatic synthesis of PGPR. However, it was experimentally demonstrated that, under the assayed experimental conditions, these lipases are not capable of catalysing the production of estolides with an acid value lower than 50 mg KOH/g (data not shown), so that, the noticeable decrease of the acid value observed in the above described experiments can be attributed mainly to the esterification reaction between polyricinoleic acid and polyglycerol. In case of the reactions catalysed by the remaining lipases tested, there is no doubt about the cause of the decrease of acid value, because they are 1,3-specific and cannot act on hydroxy fatty acids [18]. Table 7. Selection of lipases based on kinetic and economic aspects

Enzyme1

Source

4 5

Mucor miehei Pseudomonas cepacia P. fluorescens (36 U/mg solid) Rhizopus arrhizus Aspergillus oryzae Mucor javanicus P. fluorescens(309 U/mg solid) Rhizomucor miehei Chromobacterium viscosum Pseudomonas sp. (2324 U/mg solid) Pseudomonas sp. Type B (256 U/mg solid) Rhizopus oryzae

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6 7 10 12 14 15 17 18 19 20

Final AV (mg KOH/g) after 7 days 8.0 8.1

Δ AV2

Enzyme cost3 (€)

€/unit AV4

34.0 33.9

46.8 35.7

1.4 1.1

7.2

34.8

35.3

1.0

11.0 11.4 8.7

31.0 30.6 33.3

22.9 85.2 22.0

0.7 2.8 0.7

9.6

32.4

110.5

3.4

9.3 7.1

32.7 34.9

27.4 43.4

0.8 1.2

7.7

34.3

44.6

1.3

7.6

34.4

43.0

1.2

13.9

28.1

1.8

0.06

1

Enzyme identification numbers are the same that those used in Table 6. Calculated as the difference between initial AV (42.0 mg KOH/g) and final AV (3rd column). 3 Estimated from commercial price lists. 4 Calculated as column 5 divided by column 4. 2

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On the other hand, it may surprise that Mucor javanicus and Rhizopus sp. lipases (1, 3-specific) performed so well. If polyglycerol-3 is a linear molecule only two of the five hydroxyl groups available as acyl acceptor groups are primary, and the acid value reached when these lipases are used indicates that more than two hydroxyl groups has been esterified. This fact can be explained if condensation of glycerol takes place between secondary-primary or secondary-secondary hydroxyl groups. In that case more than two primary hydroxyl groups may remain available as acyl acceptor groups. As can be seen in Figure 16, satisfactory results were obtained when the twelve mentioned lipases were used to catalyse the production of PGPR and some graphs are indistinguishable. Table 7 shows the acid values reached after 7 days of reaction, which permits a better comparison of the obtained results. It can be observed that the lowest acid values were reached when lipases from Pseudomonas (3 enzymes) and Chromobacterium viscosum were used. However, some of the lipases used in the present work are very expensive, which is an aspect that should be carefully considered if the long-term purpose is to develop an industrial procedure for PGPR production. Therefore, in order to finally choose one or more of these lipases, we took into account not only kinetic aspects (reaction rates and final acid value of the reaction mixture) but also the cost of the procedure. In order to evaluate this economic aspect of the enzymatic biosynthesis of PGPR and because lipase is the most expensive material involved in the reaction, the cost of biocatalysts that cause a decrease of one unit of the acid value was calculated and the results are showed in the last column of Table 7. It can be observed that the cheapest procedures were those catalysed by lipases from the fungi Rhizopus oryzae, Rhizopus arrhizus, Mucor javanicus, Rhizomucor miehei and Rhizopus niveus, with which the decrease of one unit in acid value costs less than 1 €. These results, together with those shown in Figure 16, led us to select lipases from Rhizopus oryzae, Rhizopus arrhizus and Mucor javanicus for further experiments.

3.2.1.2. Immobilisation of the Selected Lipases Although the above selected lipases are not very expensive, to develop an industrial procedure for PGPR synthesis it is desirable to use immobilised enzymes because of its wellknown advantages: continuous operation of reactors and/or the reusability of the immobilised enzymes, both of which diminish operational costs. Therefore, the three chosen lipases were immobilised by physical adsorption onto an anion exchange resin (Lewatit MonoPlus MP 64). As described in Section 3.1.2, the authors have optimized the immobilisation process of Candida rugosa lipase and, as a preliminary attempt the same technique was used in this work in order to compare the behaviour of these three lipases. In further studies the immobilisation process should be optimized. Thus, three immobilised derivatives were prepared following the method described in Section 2.2.2 and the results are shown in Table 8, where the protein content of the commercial lipases, the immobilisation yields and the enzyme loadings of all the immobilised derivatives are summarized, all data being based on the protein concentration values provided by Lowry‘s method [16]. It should be mentioned here that the protein content of the three commercial preparations was quite low, although that of the lipase from Rhizopus arrhizus was slightly higher than the others. However, the percentage of immobilised protein obtained with this lipase was approximately half that obtained with the other two lipases, and so the enzyme loading factor of this immobilised derivative was the lowest (8.59 mg E/g support).

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The highest immobilisation yield was achieved when the lipase from Rhizopus oryzae was adsorbed; in this case an immobilised derivative with adequate enzyme content was obtained despite of the low Lowry protein content of the commercial enzyme. The immobilised derivative of lipase from Mucor javanicus had the higher enzyme loading, 14.11 mg E/g support. The above results did not differ sufficiently to permit us to decide at this stage which of the three lipases should be selected. Therefore the immobilised derivatives were tested for activity, using them to catalyse the synthesis of PGPR following the procedure described in Section 2.2.4.2. Figure 17 shows the variation of the acid value of the reaction mixtures with time. As can be seen, all the immobilised derivatives showed their ability to catalyse the esterification between polyricinoleic acid and polyglycerol-3. The use of the lipase from Mucor javanicus should not be totally discarded because reasonably good results were obtained when it was used as biocatalyst and a PGPR with an acid value of 13 mg KOH/g was reached at the end of the experiment. Table 8. Coupling parameters for the immobilisation of lipases onto Lewatit MonoPlus MP 64

Lipase source Mucor javanicus Rhizopus arrhizus Rhizopus oryzae

Protein content of the commercial lipase (%) 22.07

Immobilisation yield (%)

Enzyme loading (mg E/g support)

63.90

14.11

26.35

32.59

8.59

19.80

65.03

12.88

AV (mg KOH/g)

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50 40 30 20 10 0 0

50

100

150

200

Time (h) Figure 17. Evolution of acid value with time for the PGPR synthesis catalysed by immobilised lipases from different sources. (♦) Mucor javanicus, (▲) Rhizopus arrhizus and (×) Rhizopus oryzae.

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The highest reaction rates were achieved when lipases from Rhizopus arrhizus and Rhizopus oryzae were used and, in these cases, PGPRs with lower acid values were produced. Comparing these results with those obtained with the soluble enzymes, it can be observed that the acid values reached with the immobilised derivatives (10.42 mg KOH/g with lipase from Rhizopus arrhizus and 9.22 mg KOH/g with lipase from Rhizopus oryzae) were similar to those obtained with the soluble lipases (11.04 mg KOH/g and 13.94 mg KOH/g, respectively), even though the amounts of soluble enzymes added to the reactors (500 mg in both cases) were higher than those used in the experiments with immobilised enzymes (42.95 mg lipase from Rhizopus arrhizus and 64.4 mg lipase from Rhizopus oryzae). These results suggest that immobilisation had a beneficial effect on the activity and stability of both lipases. Lipase from Rhizopus arrhizus was selected for further studies because lower amount of enzyme is required to achieve the same final acid value.

3.2.2. Synthesis of Polyglycerol Polyricinoleate with Rhizopus arrhizus Lipase After a detailed literature search, no references were found, which might indicate, even approximately, the experimental conditions in which the reaction should be carried out. Therefore, it was necessary to start the study of the enzymatic synthesis of PGPR by establishing the optimal experimental conditions. Polyricinoleic acid (PR), which is used as substrate in all the experiments described in this section, was polymerised in our laboratory until it reached an acid number lower than 50 mg KOH/g. At this acid number value, the average length of the PR chains is four, which is considered as optimum for food use by most authors [3, 13]. Among the commercially available polyglycerols, Polyglycerol-3 from Solvay (PR-3) was considered the most appropriate because it produces a high performance PGPR [34]. As described in Section 1, the European Commission Directive 2008/84/EC [14] establishes an acid value (AV) lower than 6 mg KOH/g for PGPR and a hydroxyl value (HV) of between 80 and 100 mg KOH/g. HV is a measurement of the free hydroxyl groups and any reduction is concomitant with a decrease in AV because one of each is consumed when an ester linkage is formed [35]. When the PR/PG-3 mass ratio used is lower than 12 both requirements cannot be fulfilled because, even if all the acid groups react and AV is close to zero, too many hydroxyl groups remain unreacted (HV > 100 mg KOH/g). On the other hand when the PR/PG-3 ratio is too high, the final product will contain too many acid groups or too few hydroxyl groups. Therefore, the substrate mass ratio (PR/PG-3) in all the experiments was maintained constant at a value of 15, which means that three of the five hydroxyl groups of the polyglycerol could be esterified. On the other hand, it is important to underline the dramatic influence of relative humidity on the equilibrium of this esterification process. To avoid this effect, all the experiments used to optimize an individual variable were carried out simultaneously, in an air open tank reactor when the relative humidity was 40-50 %. 3.2.2.1. Influence of Initial Water Content In order to establish the optimal initial amount of water in the reactor, a set of experiments was carried out with 500 mg lipase, varying the volume of water used to dissolve

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the enzyme from 0.75 to 10 ml of water. Since similar results were obtained in all the experiments (data not shown), 5 ml of distilled water was used in further studies when the amount of enzyme added was 500 mg.

3.2.2.2. Influence of Temperature The effect of the temperature on the reaction course was investigated and three experiments were carried out at 40, 50 and 60ºC. Figure 18 shows the evolution of the acid value of the reaction mixture with reaction time. The adverse effect observed on the reaction when the experiment was carried out at the highest temperature, can be attributed to a thermal deactivation of the lipase, since it is well known that most proteins tend to denaturalize at temperatures above 50ºC. The most common cause for the inactivation of enzymes at high temperatures is loss of the native, catalytically active conformation, i.e. thermodenaturation. Because similar acid values were achieved at 40 and 50ºC, the first one was used in the following experiments, especially because 40ºC has been described as the optimum temperature for Rhizopus arrhizus lipase [36]. 3.2.2.3. Influence of the Amount of Enzyme For the synthesis of polyglycerol polyricinoleate, the influence of different amounts of added lipase on the esterification reaction was studied in five experiments using 50, 100, 200, 500 and 1000 mg of lipase. As can be seen in Figure 19, a significant improvement in the reaction extension was obtained when the amount of added enzyme was increased from 50 mg (minimum AV=22.02 mg KOH/g) to 500 mg (minimum AV=17.76 mg KOH/g). However, no additional enhancement was observed when 1000 mg of lipase was added to the reaction mixture (AV=17.69 mg KOH/g). As a consequence, 500 mg lipase was used in further experiments.

AV (mg KOH/g)

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50 40 30 20 10 0 0

50

100

150

Time (h) Figure 18. Influence of temperature on PGPR production using free Rhizopus arrhizus lipase. (  ) 40ºC, (  ) 50ºC and (  ) 60ºC.

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AV (mg KOH/g)

50 40 30 20 10 0 0

50

100

150

200

Time (h)

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Figure 19. Influence of the amount of lipase on PGPR production. (  ) 50 mg, (  ) 100 mg, (  ) 250 mg, (  ) 500 mg and (  ) 1000 mg of Rhizopus arrhizus lipase dissolved in 5 ml of distilled water.

3.2.2.4. Lipase Immobilisation and Comparison with Free Enzyme All the experiments mentioned until now were carried out adding the native enzyme to the reaction medium. It is clear that the acid value of the synthesised PGPG did not fulfil the requirements demanded by the European Commission Directive 2008/84/EC [14]. Moreover, as shown in Figure 19, when the amount of lipase added to the reactor was increased, the reaction progressed faster and a lower acid value was reached. The differences between the acid values reached led us to think that these were not equilibrium values but that the reaction stopped because the enzyme was no longer active. Taking into consideration the potential benefits from the use of immobilised enzymes (high activity and stability), the possibility of immobilising Rhizopus arrhizus lipase by physical adsorption onto an ion exchange resin (Lewatit MonoPlus MP 64) was studied. The immobilisation method is illustrated in Section 2.2.2. As we previously mentioned (Section 3.2.1.2), when Rhizopus arrhizus lipase was immobilised in the conditions considered as optimum for Candida rugosa lipase, an immobilised derivative containing 8.6 mg of protein per g of support was obtained with an immobilisation yield of 32.6 %. A 5 g sample of this derivative (containing 43 mg of protein) were used as catalyst for the synthesis of PGPR and, in order to compare the behaviour of immobilised and free lipases, a similar synthesis was carried out with the same amount of protein (since the commercial preparation from Fluka contains 26.3% of protein, 163.5 mg of commercial free enzyme were added). Figure 20 shows the results of these experiments. Two conclusions can be reached for the curves observed. First, the immobilisation provokes a moderate loss of lipase activity since the initial reaction rate (24 h) is lower when immobilised lipase is used. Secondly, immobilisation seems to stabilize the lipase activity: after 168 h the condensation process still continues with immobilised lipase, while the reaction does not continue beyond 50 h when free lipase is used.

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AV (mg KOH/g)

50 40 30 20 10 0 0

50

100

150

200

Time (h)

3.2.2.5. Reuse of the Immobilised Derivative Immobilisation provides an attractive opportunity for the multiple use of the enzyme. In order to establish the reusability of our immobilised derivative (lipase), successive polymerisation reactions were carried out, examining the evolution of AV in several consecutive experiments. Not many immobilised enzymes exhibit such a good reusability as that shown in Figure 21 by Rhizopus arrhizus lipase. Three consecutive processes yielded an almost identical acid values after seven days of operation. Moreover, the immobilised derivative could be easily removed from the reaction medium, conventional filtration through a sinterized glass filter and gravity force being sufficient to successfully separate the derivative from the viscous product, PGPR. 50 40

AV (mgKOH/g)

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Figure 20. Influence of enzyme immobilisation on PGPR production. (  ) 163 mg of free lipase dissolved in 5 ml of distilled water or (  ) 5 g of the immobilised derivative and the amount of water soaked in the support.

30 20 10 0 0

50

100

150

Time (h)

Figure 21. Reusability of the immobilised derivative in PGPR production. (  ) First use, ( ) second use and (  ) third use. Lipase: Functions, Synthesis and Role in Disease : Functions, Synthesis and Role in Disease, Nova Science Publishers, Incorporated, 2012.

Solvent-Free Biocatalytic Synthesis of Polyglycerol Polyricinoleate ...

107

50

AV (mg KOH/g)

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3.2.2.6. Obtaining PGPR in the Vacuum Reactor It was mentioned above that the European Commission Directive 2008/84/EC [14] establishes as a requirement for PGPR an acid value lower than 6 mg KOH/g. In all the experiments described until now the final acid value was far from this objective, which means that the equilibrium has to be shifted towards the esterification pathway. This can be done by using a more anhydrous medium. On the other hand, the crucial importance of the amount of water in the reaction medium is illustrated by the poor reproducibility of the processes taking place in open reactors, since this parameter is heavily influenced by seasonality and weather conditions. This is particularly important in our case, when the final purpose is the production of the additive on an industrial scale, which requires rigorous standardisation. For this reason, a high performance reactor was tested for PGPR production. The reactor is thermostated, it can work in a wide range of pressures, and is also able to mix to the reaction medium in accordance with its high viscosity. In this reactor the amount of water can be manipulated through the pressure and the entry of dry nitrogen, making it independent on laboratory conditions. Figure 22 shows the results of PGPR synthesis using immobilised lipase in the high performance reactor. This reactor (described in Section 2.2.4), provides a controlled atmosphere that facilitates the adjusting of the water content in the reactor medium. The pressure was set at 160 mm Hg and 90 l/h dry nitrogen was continuously passed through the reaction mixture. In these conditions, the water content in the reaction medium was stabilised at around 2000 ppm (Karl-Fisher) after 10 h (a totally anhydrous medium would lead to enzyme inactivation). In these conditions, the objective of the European Commission Directive was attained after 100 h (AV = 5.9 mg KOH/g) but even lower values can be reached at longer times (after 125 h the AV was 4.9 mg KOH/g). Although Figure 22 only depicts one curve corresponding to the vacuum reactor, several experiments were carried out in identical conditions and the same results were obtained.

40 30 20 10 0 0

40

80

120

160

Time (h) Figure 22. Influence of the reactor device on the production of PGPR using immobilised Rhizopus arrhizus lipase. (  ) Open air reactor and (  ) vacuum reactor.

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Figure 22 also reveals the enormous difference between PGPR biosynthesis in open air and vacuum reactors. Both esterification processes were made with the same amount of substrate, immobilised derivative and initial amount of water. In the atmospheric reactor the target was not reached even after one week, whereas in the vacuum reactor PGPR was ready after 100 h.

CONCLUSION

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The biocatalytic synthesis of polyglycerol polyricinoleate (E-476) from polyglycerol and ricinoleic acid is described in detail in this work. The process is carried out in two steps: the first one the production of polyricinoleic acid is successfully catalysed by Candida rugosa lipase and, the second one, the esterification of polyricinoleic acid with polyglycerol is catalysed by Rhizopus arrhizus lipase. The process is carried out as a solvent free system, a feature that enhances the greenchemistry reputation that enzymatic processes already have. Moreover, the biosynthesis is catalysed by immobilised derivatives with excellent reusability, which makes the process even more advantageous from the economic and environmental point of view. When the biosynthesis of PGPR is carried out in an open air reactor, atmospheric humidity does not allow a sufficient degree of esterification, yielding a product with an acid value that clearly exceeds that established by the European Commission Directive for PGPR. However, when PGPR is obtained in a controlled atmosphere with low humidity, the esterification equilibrium swifts towards the synthetic pathway, yielding a product that fulfils all the specifications of the European Commission.

ACKNOWLEDGMENTS S. Ortega is beneficiary of a pre-doctoral scholarship from Fundación Séneca of Murcia.

REFERENCES [1] [2] [3] [4]

[5]

Wilson, R.; Van Schie, B. J. & Howes, D. (1998). Overview of the preparation, use and biological studies on polyglycerol polyricinoleate. Food Chem. Toxicol., 36, 711-718. Denecke, P.; Börner, G. & Allmen, V. V. (1981). Method of preparing polyglycerol polyricinoleic fatty acid esters. UK Patent Application 2,073,232 A. Bódalo, A.; Bastida, J.; Máximo, M. F.; Montiel, M. C. & Murcia, M. D. (2005). Enzymatic biosynthesis of ricinoleic acid estolides. Biochem. Eng. J., 26, 155-158. Bódalo, A.; Bastida, J.; Máximo, M. F.; Montiel, M. C.; Gómez, M. & Murcia, M. D. (2008). Production of ricinoleic acid estolide with free and immobilised lipase from Candida rugosa. Biochem. Eng. J., 39, 450-456. Bódalo, A.; Bastida, J.; Máximo, M. F.; Montiel, M. C.; Murcia, M. D. & Ortega, S. (2009). Influence of the operating conditions on lipase-catalysed synthesis of ricinoleic acid estolides in solvent-free system. Biochem. Eng. J., 44, 214-219.

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Solvent-Free Biocatalytic Synthesis of Polyglycerol Polyricinoleate ... [6]

[7]

[8]

[9]

[10]

[11]

[12] [13] [14]

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

[15]

[16] [17] [18] [19] [20]

[21] [22] [23]

109

Bódalo, A.; Bastida, J.; Máximo, M. F.; Montiel, M. C.; Gómez, M. & Ortega, S. (2009). Screening and selection of lipases for the enzymatic production of polyglycerol polyricinoleate. Biochem. Eng. J., 46, 217-222. Gómez, J. L.; Bastida, J.; Máximo, M. F.; Montiel, M. C.; Murcia, M. D. & Ortega, S. (2011). Solvent-free polyglycerol polyricinoleate synthesis mediated by lipase from Rhizopus arrhizus. Biochem. Eng. J., 54, 111–116. Santos, J. C.; Nunes, G. F. M.; Moreira, A. B. R.; Perez V. H. & Castro, H. F. (2007). Characterization of Candida rugosa lipase immobilised on poly (Nmethylolacrylamide) and its application in butyl butyrate synthesis. Chem. Eng. Technol., 30, 1255-1261. Guo, L.; Zhang, Z.; Zhu, Y.; Li, J. & Xie, Z. (2008). Synthesis of polysiloxanepolyester copolymer by lipase-catalysed polycondensation. J. Appl. Polym. Sci., 108, 1901-1907. Dang, H. T.; Obiri, O. & Hayes, D. G. (2005). Feed batch addition of saccharide during saccharide-fatty acid esterification catalysed by immobilised lipase: time course, water activity and kinetic model. J. Am. Oil Chem. Soc., 82, 487-493. Goldberg, M.; Thomas, D. & Legoy, M. D. (1990). The control of lipase-catalysed transesterification and esterification rates. Effects of substrate polarity, water activity and water molecules on enzyme activity. Eur. J. Biochem., 190, 603–609. Yahya, A. R. M.; Anderson, W. A. & Moo-Young, M. (1998). Ester synthesis in lipase catalysed reactions. Enzyme Microb. Technol., 23, 438-450. Yoshida, Y.; Kawase, M.; Yamaguchi, C. & Yamane, T. (1997). Enzymatic synthesis of estolides by a bioreactor. J. Am. Oil Chem. Soc., 74, 261–267. Commission Directive 98/86/EC of 11 November 1998, published in Official Journal of the European Communities L334/1 of 9 December 1998. Bódalo, A.; Gómez, E.; Gómez, J. L.; Bastida, J.; Máximo, M. F. & Díaz, F. (1991). A comparison of different methods of β-galactosidase immobilisation. Process Biochem., 26, 349–353. Hartree, E. F. (1973). Protein determination and improved modification of the Lowry‘s method which gives a linear photometric response. Anal. Biochem., 42, 422–427. ASTM D974-06. Standard test method for acid and base number by color indicator titration. Hayes, D. G. (1996). The catalytic activity of lipases toward hydroxy fatty acids. A review. J. Am. Oil Chem. Soc., 73, 543–549. Yoshida, Y.; et al. (1993). Production of estolide from ricinoleic acid. Japanese Patent, JP521878 (in Japanese). Gómez, J. L.; Bódalo, A.; Gómez, E.; Bastida, J.; Hidalgo, A. M. & Gómez, M. (2006). Immobilisation of peroxidase on glass beads: an improved alternative for phenol removal. Enzyme Microb. Technol., 39, 2016– 2022. Yokomichi, H.; Yasumasi, T.; Nakamura, K. & Kawahara, Y. (1988). Immobilised enzyme and esterification and interesterification therewith. EPO Patent EP0320132. Gitlesen, T.; Bauer, M. & Adlerereutz, P. (1997). Adsorption of lipase on polypropylene powder. Biochim. Biophys. Acta, 1345, 188–196. Yesiloglu, Y. (2005). Utilization of bentonite as a support material for immobilisation of Candida rugosa lipase. Process Biochem., 40, 2155–2159.

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[24] Chiou, S. H. & Wu, W. T. (2004). Immobilisation of Candida rugosa lipase on chitosan with activation of the hydroxyl groups. Biomaterials, 25, 197–204. [25] Domínguez de María, P.; Xenakis, A.; Stamatis, H. & Sinisterra, J. V. (2004). Unexpected reaction profile observed in the synthesis of propyl laurate when using Candida rugosa lipases immobilised in microemulsions based organogels. Biotechnol. Lett., 26, 1517–1520. [26] Ye, P.; Xu, Z. K.; Wang, Z. G.; Wu, J.; Deng, H. T. & Seta, P. (2005). Comparison of hydrolytic activities in aqueous and organic media for lipases immobilised on poly(acrylonitrile-co-maleic acid) ultrafiltration hollowfiber membrane. J. Mol. Catal. B Enzym., 32, 115–121. [27] O‘Connell, P. J. & Varley, J. (2001). Immobilisation of Candida rugosa lipase on colloidal gas aphrons (CGAs). Biotechnol. Bioeng., 74, 264–269. [28] Panzavolta, F.; Sore, S.; D‘Amato, R.; Palocci, C.; Cernia, E. & Russo, M.V. (2005). Acetylenic polymers as new immobilisation matrices for lipolytic enzymes. J. Mol. Catal. B Enzym., 32, 67–76. [29] Kelly, A. R. & Hayes, D. G. (2006). Lipase-catalysed synthesis of polyhydric alcoholpoly(ricinoleic acid) ester star polymers. J. Appl. Polym. Sci., 101, 1646–1656. [30] Klibanov, A. M. (1989). Enzymatic catalysis in anhydrous organic solvents. Trends in Biochemical Sciences, 14, 141-144. [31] Basheer, S.; Mogi, K. & Nakajima, M. (1995). Surfactant-modified lipase for the catalysis of the interesterification of triglycerides and fatty acids. Biotechnol. Bioeng., 45, 187–195. [32] Hayes, D. G. & Kleiman, R. (1995). Lipase-catalysed synthesis and properties of estolides and their esters. J. Am. Oil Chem. Soc., 72, 1309–1316. [33] Hayes, D. G. & Kleiman, R. (1996). Lipase-catalysed synthesis of lesquerolic acid wax and diol esters and their properties. J. Am. Oil Chem. Soc., 73, 1385-1392. [34] Polyglycerols in Food Applications. Application Data Sheet. http://www.solvaypolyglycerol.com/docroot/glycerol/static_files/attachments/polyglyce rols_for_food.pdf [35] Committee on Food Chemicals Codex, Food and Nutrition Board, Institute of Medicine. (2004). Food Chemicals Codex 5th ed. Washington: The National Academies Press. [36] Méndez, J. J.; Canela, R. & Torres, M. (2009). Kinetic study of palmitic acid esterification catalysed by Rhizopus oryzae resting cells. Acta Biol. Colomb., 14, 161172.

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In: Lipase Editors: Hamdi Sassi and Sofien Cannamela

ISBN 978-1-62081-366-9 © 2012 Nova Science Publishers, Inc.

Chapter IV

FUNGAL LIPASES: VERSATILE TOOLS FOR BIOCATALYSIS Selmene Ouertani1 and Habib Horchani2, 1

Laboratoire de Biochimie et de Génie Enzymatique des Lipases, Tunisie LOEX/CUO-recherche, Centre hospitalier affilié universitaire de Québec and Département d‘opthtalmologie, Faculté de Médecine, Université Laval, Québec, Canada 2

ABSTRACT

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Lipases are ubiquitous enzymes which catalyze the hydrolysis of triacylglycerols to glycerols and free fatty acids. Under certain experimental conditions, such as in the absence of water, they are capable of reversing the reaction. The reverse reaction leads to esterification and formation of Mono-, Di-, Tri-glycerides from fatty acids and glycerol. Due to their very interesting characteristics like high enantioselectivity and regioselectivity and stability in organic solvents, lipases have gained in the last decade much more importance in industrial applications. Among these biocatalysts, fungal lipases represent an important family due to their low cost of extraction, thermal and pH stability, substrate specificity and activity in organic solvents. In fact, many fungal lipases have been produced, purified and biochemically characterized since the middle of the last century. The chief producers of commercial lipases are Aspergillus niger, Candida cylindracea, Humicola lanuginosa, Mucor miehei, Rhizopus arrhizus, Rhizopus delemar, Rhizopus japonicus, Rhizopus niveus and Rhizopus oryzae. These lipases are usually used in a variety of biotechnological fields such as food and dairy (cheese ripening, flavor development), detergent, pharmaceutical, agrochemical (insecticide, pesticide) and oleochemical (fat and oil hydrolysis, biosurfactant synthesis, polymer synthesis) industries…



Corresponding author: Habib Horchani, LOEX/CUO-recherche, Centre hospitalier affilié universitaire de Québec and Département d‘opthtalmologie, Faculté de Médecine, Université Laval, Québec, Canada, [email protected].

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Selmene Ouertani and Habib Horchani

INTRODUCTION

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Enzymes have found great uses in several industries such as food, dairy, pharmaceutical, detergent, textile, animal feed, and cosmetics. The number of enzymes commercially available and the range of applications are gradually increasing [1]. There are many reasons for the growing interest in enzyme-mediated reactions compared to chemical processes, including high degree of specificity, mild reaction conditions, decrease in side reactions, and simplicity of post-recuperation. Furthermore, enzyme-mediated processes are energy saving and reduce the extent of thermal degradation [2,3]. Among all enzymes, lipases (triacylglycerol acylhydrolase, EC 3.1.1.3), a class of enzymes that catalyze the hydrolysis of the ester bonds in triacylglycerols, are gaining more importance. Lipases catalyze three types of reversible reactions (Figure 1). They catalyze hydrolysis in an aqueous system, but also esterification (non conventional medium) in a microaqueous system, where water content is very low. Transesterification is categorized into four subclasses according to the chemical species which react with the ester (Figure 1). Alcoholysis is the reaction with an ester and an alcohol, while acidolysis is the one with an ester and an acid. Interesterification is a reaction between two different esters, where alcohol and acid moiety are swapped. In aminolysis, an ester is reacted with an amine, generating an amide and an alcohol. The great interest in lipases is mainly owing to their properties in terms of enantioselectivity, regioselectivity and broad substrate specificity [4,5]. The use of lipases as biocatalysts for the production of biomolecules has many more potential benefits for future developments besides their specificity. The most important merits are ● ● ● ● ●

Efficacy of lipases under mild reaction conditions, Utility in ―natural‖ reaction systems and products, Reduction of environmental pollution, Availability of lipases from a wide range of sources, Ability to improve lipases by genetic engineering,

For these reasons, many nutritional and functional molecules with high added value have been produced enzymatically and a lot of these studies have been published in the past twenty years. Although lipases are of widespread occurrence throughout the Earth‘s flora and fauna, they are found more abundantly in microbial flora comprising bacteria, fungi and yeast [4,6,7]. Furthermore, microbial lipases, compared with the other sources of lipases, are commercially significant because of low production cost, greater stability and wider availability than plant and animal lipases. In fact, Most of the industrial microbial lipase is derived from fungi and bacteria [8]. Fungal lipases have gained special industrial attention due to their stability, selectivity, and broad substrate specificity [9-11]. The ability of lipases to perform very specific chemical transformation (biotransformation) has made them increasingly popular in the food, detergent, chemical, and pharmaceutical industries [12,13]. The present chapter will concentrate on the lipases produced by fungi and will focus on their biotechnological applications.

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113

Figure 1. Hydrolyse, esterification and transesterification reactions catalysed by lipases.

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ESSENTIAL CHARACTERISTICS OF FUNGAL LIPASES Fungal lipases have benefits over bacterial lipases due to their low cost of extraction, thermal and pH stability, substrate specificity and activity in organic solvents. Lipase producers are widespread in the fungal kingdom. Some of the major lipase-producing fungi are of the genera Mucor, Rhizopus, Geotrichum, Rhizomucor, Aspergillus, Humicola, Candida, and Penicillium [14-16]. Lipase production by fungi varies according to the strain, the composition of the growth medium (the kind of carbon and nitrogen sources used), cultivation conditions, temperature and pH [17]. The fungal lipase have optimum pH range from 4 to 8 [18,19] with some exceptions, like the Aspergillus niger NCIM 1207 lipase [20] and the Aspergillus Carneus lipase [21] which showed an optimum pH of 2.5 and 9, respectively. Most of the fungi have an optimum of lipase activity at a temperature between 25-30°C except some thermophilic fungi which are more active at high temperature (45-75°C) [22-24]. One can also note that the production of the extracellular and cell bound lipases were reported to depend on the Carbon and Nitrogen composition of the medium [24]. Table 1 highlights the biochemical properties of some fungal lipases recently studied.

BIOTECHNOLOGICAL APPLICATIONS OF FUNGAL LIPASES There has been a tremendous increase in the significance of the biotechnological applications of fungal lipases since the middle of the last century [35,36], due to the versatile

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catalytic behavior of lipases [37]. Lipases form an integral part of the industries ranging from food [38], pharmaceuticals [39] and detergents [40] to oleo-chemicals [13], tea industries [41], agriculture, cosmetics, leather and in several bioremediation processes. Fungal Lipases are used in two distinct fashions, they are used as biological catalysts to manufacture other products (such as food ingredients) and by their application as such. Because of their high performance, some fungal lipases are commercialized for one specific application whereas others can be used in different industrial fields. Table 2 highlights some commercial fungal lipases from different suppliers.

MW (kDa)

pH optimum

Optimum Temperature (°C)

Specific activity (U/mg)

Other Characteristics

Rhizopus homothallicus LipA

29,5

7,5

40

8600

- Half life time at 50 ° C is 0.4 h.

LipB

29,5

7,5

30

10700 (TC8)

Penicilllium camembertii Thom PG-3

28

6,4

48

nd

Rhizopus oryzae

32

8

37

Rhizopus oryzae. W

29

8

37

10000 (olive oil) 3500 (olive oil)

Penicillium aurantiogriseum

28

8

60

17.8 (TC4)

Aureobasidium pullulans HN2.3 Antrodia cinnamomea BCRC 35396 Aspergillus niger NCIM 1207

63,5

8,5

35

60

8

45

32,2

2,5

50

88,6 (p-NPL) 187,5 (p-NPP) 1373,13 (p-NPP)

Aspergillus carneus

27

9

37

502 (olive oil)

Aspergillus niger

32

3-5

25-35

Aspergillus niger MZKI Al 16

65

7

45

627 (Tricaprylin) nd

Aspergillus oryzae

21,6

9

55

694 (pNPButyrate)

- Half life time at 50 ° C is 0.4 h. -1,3 positional specificity.

References

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Table 1. Biochemical properties of some fungal lipases

[25]

[26]

- Specificity towards long chain TG. - Doesn‘t show the mechanism of interfacial activation. -Stable in organic solvents

[27]

-Inhibited by Hg2+, Fe2+ and Zn2+. -Alkaline lipase.

[30]

-Acidic lipase. -Specific for 3position in the ester bond. -1,3 positional specificity. - Alkaline lipase. -Thermostable -Sn-1 specificity. -Specificity towards short chain of TG. -1,3 positional specificity. -Degradation of bioplastics.

[20]

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[28]

[29]

[31]

[21]

[32] [33]

[34]

Fungal Lipase

115

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Table 2. Some commercially available fungal lipases, their sources and industrial applications Source

Application

Humicola lanuginose Candida cylindracea

Detergent additive Food processing

Rhizomucor miehei Thermomyces lanuginosus

Food processing

Aspergillus niger

Food processing

Rhizopus oryzea

Food processing, oleochemistry

Mucor miehei

Food processing

Penicillium camemberti Penicillium sp.

Food processing, oleochemistry Food processing

Candida antarctica A/B

Organic synthesis

Detergent additive

Commercial name Lipolase TM

Company

References

Novo Nordisk

[35]

ChiroCLEC-CR Lipase AY Lipase MY, Lipase OF-360 Chirazyme® L-3

Atlus Biologics Amano Meito Sangyo

[13, 24]

Lipomod™ 34PL034P Palatase® Lipolase®, Lipolase® Ultra, Lipo Prime™, Lipex® Lipase A ―Amano‖ 6 Lypolyve AN Lipase F-AP15 Lipomod™ 627P-L627P Piccnate Lipase G ―Amano‖ 50 Lipomod™ 621P-L621 Chirazyme®L-5 SP526 Chirazyme®L-2 SP 525 or Novozym 435b

Boehringer Mannheim Biocatalysts Novozymes

[13]

Novozymes

[13]

Amano

[13]

Lyven Amano Biocatalysts

[13]

Gist-Brocades Novo Nordisk Amano

[13]

Biocatalysts

[13]

Boehringer Mannheim Nova-nordisk Boehringer Mannheim Nova-nordisk

[24]

[13]

1. Food Industry Lipases have become an integral part of the modern food industry [42]. Industrial lipases, especially fungal ones, are commonly used in the production of a variety of products, ranging from baked foods and oleochemical industry to dairy enrichment [38]. Some unique

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properties of fungal lipase such as their specificity, temperature, pH dependency and nontoxic nature lead to their major contribution in the different areas of food processing industries. Fungal lipases have also been used in the production of sugar fatty acid esters which are very useful in different industrial areas. For example, Penicillium camembertii lipase was used for the production of glycerol-glycolipid from a mixture containing glycosidases and fatty acid [43]. Silica granulated Thermomyces. lanuginosa lipase served as feasible biocatalyst for the synthesis of acylsucroses or other carbohydrates esters in the polar or slightly polar solvents [44]. Lipases from Thermomyces. lanuginosus and Candida antarctica B were used as a potentially reliable catalysts for sugar ester production [45]. In confectionary, 1,3-regioselectivity of lipases was exploited in the process development of a fat production containing high concentration of 1,3-disteraroyl-2-monoloein [46]. This fat could be used as a substitute for shea stearine in the formulation of cocoa butter equivalents [38]. The interesterification by 1,3-regio specific lipases have been also used to enrich the palm-oil fractions in to 1, (3) polmitoyl, 2-oleoyl, 3(1) stearoyl glycerol and 1(3) steraroyl, 2-oleoyl 3(1) stearoylglycerol, which found immense application as the confectionary fats (Unilever, 1980) [47].

Fats and Oil Industry Fats and oil modification is one of the prime areas in food processing industry that demand novel economic and green technologies [48]. Fats and oils are important constituents of foods. Lipases allow us to modify the properties of lipids by altering the position of fatty acid chains in the glyceride and replacing one or more of these with new ones. In this way, a relatively inexpensive and less desirable lipid can be modified to a higher value fat [8]. Many authors have reported the use of different fungal lipases for the modification of oils and the upgrade of cheap oils and fats to synthesize nutritionally important lipids [50-53]. The Rhizopus japonicus lipase has been used to produce hard butter suitable for chocolate manufacture by interesterification of palm oil with methyl stearate [49]. It has been also demonstrated by Safari and Kermasha [50] that, among several commercial enzymes, lipase from Rhizopus niveus shows an interesting potential for the production of interesterified butter fat with an increased proportion of oleic acid at the sn-2 position [50]. The interesterification of olive oil with palmitic acid catalyzed by Rhizopus delemar lipase was investigated in phospholipids microemulsion systems by Komatsu et al. using soybean lecithin as the amphiphilic component [51]. Rhizomucor miehei lipase and lipozyme IM20 were also used for the modification of tallow by interesterification of its stearine fractions with liquid oils (the sunflower, soybean, rice bran,…) and acidolysis of its olein fractions with karanja (Pongamia glabra) stearine. The products formed, having a slip melting point and solid fat index, were suitable for shortening the margarine fat bases and vanaspati substitute [52]. Recombinant Aspergillus oryzae lipase was used as a biocatalyst in the oils and fats industry for oil de-gumming to improve emulsifying properties [53]. Structured lipids (SL) present, depending on the molecular structures of the acyl chains, unique nutritional and functional properties [3]. Currently fungal lipases, due to their high specificity, are successfully used for the synthesis of some of these very high added value products. The SL containing palmitic, oleic, stearic and linoliec acids, resembling human milk fat, were synthesized using commercially immobilized lipozyme [54]. It has been reported by Tsuzuki [55] that Aspergillus oryzae lipase was a powerful biocatalyst to produce reducedcalorie SL by an acidolysis reaction. The results showed that the lipase promoted the

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incorporation of more than 80% of triolein by butyric acid in the dried n-hexane at 52 °C, after 72 h of reaction time. Compared to the chemical process, this method is very interesting due to a reduction in the number of byproducts [56]. Interesterification using fungal lipases can also be used to produce oils and fats containing nutritionally important polyunsaturated fatty acids (PUFAs), such as eicospentaenoic and docosahexaenoic (DHA) acids which are known to play an important role in human health [51]. One of the main applications of the lipase-catalyzed transesterification reaction is the incorporation of Docosahexaenoic Acid /Eicosapentaenoic Acid into vegetable oils [57]. Immobilized Rhizopus oryzae lipase appears as an efficient biocatalyst for DHA enrichment of a mixtures of fatty acids from sardine oil by selective esterification [58]. The Aspergillus niger lipase was one of the most effective enzymes in concentrating n-3 PUFA. The degree of hydrolysis (60%) led to an increase in the DHA content from 14% in the original oil to 34% in the residual acylglycerol [59]. A lipase from Penicillium abeanum was also used to enrich DHA in Tuna oil [60]. After the enzymatic reaction, the content of DHA in Tuna oil increased by more than 2-folds [60]. Monoglycerides (MGs), nonionic surfactants and emulsifying molecules with both hydrophilic and hydrophobic regions, are widely used in food industry [61,62]. MGs were successfully produced using fungal lipases by direct esterification of fatty acid with glycerol [63,64] or by glycerolysis [65]. For example, the Penicillium camembertii lipase immobilized on epoxy SiO2-PVA was used in the monoolein synthesis in solvent-free system with a conversion yield of about 40 % [63]. Yamaguchi and Mase have obtained a conversion yield of synthesis of monoolein of about 76% using the mono- and diacylglycerol lipase from Penicillium camembertii U-150 [64]. A high conversion yield (90 to 100%) was obtained with Novozym 435 (Candida antarctica lipase) to produce MGs from commercial oils and fats using a tetra ammonium based ionic liquid as the reaction medium [65].

Dairy Industry Fungal Lipases are extensively used in the dairy industry for the hydrolysis of milk fat. Lipases are also used to modify the fatty acid chain lengths and to enhance the flavors of various cheeses. Current applications also include the acceleration of cheese ripening and the lipolysis of butter, fat and cream [8,66]. The free fatty acids generated by the action of lipases on milk fat endow many dairy products, particularly soft cheeses, with their specific flavor characteristics. A whole range of fungal lipase preparations have been developed for the cheese manufacturing industry: Mucor meihei, Aspergillus niger and Aspergillus oryzae lipases and several others [35]. It has been reported that the addition of fungal lipases contributes strongly to the development of cheese flavors and accelerates greatly the cheese ripening process [13,35,36,38]. Peters and Nelson [67] have showed that the addition of Candida lipolytica lipase enhanced the quality of blue cheese made from unhomogenized, pasteurized milk. The Rhizopus delemar lipase, produced by the Tanabe Seiyaku Co., Japan, has been used for enhancing flavors of dairy products, such as milk, cheese, butter, etc [36]. Fungal lipases also play a crucial role in the preparation of so-called enzyme modified cheeses (EMC). It is produced when cheese is incubated in the presence of enzymes at elevated temperature in order to produce a concentrated flavor, such as dips, sauces, soups and snacks. The concentration of fat is 10 times higher in EMC than that of normal cheese [68].

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Cocoa butter contains palmitic and stearic acids and has a melting point of approximately 37°C, leading to its melting in mouth which results in a cooling sensation. In 1976, Unilever filed a patent describing a mixed hydrolysis and synthesis reaction to produce a cocoa butter substitute using an immobilized lipase. This technology was commercialized by Quest-Lodrs Croklaan, based on immobilized Rhizomucor miehei lipase, which carries out a transesterification reaction replacing palmitic acid with stearic acid to give the desired stearic–oleic– stearic triglyceride [8,69].

Flavor Production The production of esters as flavor compounds by biotechnological processes using enzymes has potential interest for the food industry. The main advantages of the enzyme catalyzed flavor generation are high selectivity or specificity of the reaction, high reaction rate even at low molar fractions, environmentally compatible and mild reaction conditions, and if viable cells are used there is a possibility to perform the multistep synthesis. Nowadays, this specific application meets the consumers demand for the natural products rather than synthetic products [47]. Several studies have described the use of fungal lipases in the synthesis of flavor compounds with great effectiveness [70-74]. For example, Immobilized lipases from Mucor miehei and Candida antarctica were used for the synthesis of short-chain flavor thioester in solvent-free medium [70]. Langrand et al. demonstrated that Aspergillus niger lipase was very useful in the synthesis of terpene alcohol esters of short fatty acids (C3–C6) [71]. Ethyl hexanoate is a typical fragrance compound of Chinese liquor and Japanese sake with an annual demand of more than 2000 kg. Whole cell lipase of Rhizopus chinensis had a much higher ability to the synthesis of ethyl hexanoate with a maximum yield of 96.5% after 72 h among ten commercial lipases studied [72]. The free Mucor miehei lipase was used as a biocatalyst to the direct esterification of citronellol and geraniol with short-chain fatty acids [73]. Hexyl butyrate synthesized by the immobilized lipase (Lipozyme IM-77) from Rhizomucor miehei was employed as flavor and fragrance in the food, beverage and pharmaceutical industry. It was synthesized by lipase catalyzed mild tranesterification of hexanol and tirbutrin. It found enormous interest as the natural flavoring compound rather artificial or synthetic [74]. Bakery Industry The use of enzymes enables bakeries to extend shelf-life of breads, enhance and control non-enzymatic browning, increase loaf volume and improve crumb structure. Lipases were primarily used to enhance the flavor content of bakery products by liberating short-chain fatty acids through esterification. Along with flavor enhancement, it also prolonged the shelf-life of most of the bakery products. Texture and softness could be improved by lipase catalysis [75]. Lipases are also effective in replacing partially or totally the emulsifiers, and to increase the volume in bread and bakery [35]. Greenough et al. reported that an artificially expressed lipase in Aspergillus oryzae was used as catalyst in the baking industry [76]. Lipases from Aspergillus niger, Rhizopus oryzae, Candida cylindracea are also used in bakery products [35].

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2. Detergent Industry The enzyme based detergents have better cleaning properties as compared to synthetic detergents. They are active at low washing temperatures and environment friendly [77]. They also improve the fabric quality and keep color bright. They are used in small quantity as compared to synthetic chemicals. The enzymes in the detergents do not lose their activity after removing stain [40]. Among these enzymes, lipases are widely used in detergents, where their function is in the removal of fatty residues and cleaning clogged drains [78]. The cleaning power of lipase (or other enzyme containing) detergents increases markedly. In certain conditions and to be a suitable additive in detergents, lipases should be thermostable, tolerant to an alkaline environment, capable of functioning in the presence of the various components of washing powder formulations and present the ability to hydrolyze fats with various compositions [2,79]. The most commercially important field of application for hydrolytic lipases is their addition to detergents, which are used mainly in household and industrial laundry and in household dishwashers [80]. Godfrey and West reported that about 1000 tones of lipases are sold every year in the area of detergents [81]. Fungal lipases function is the removal of stains from fabrics and are important components of detergent mixtures [82]. In 1988, Novo Nordisk developed a lipase produced by the fungus Humicola. Commercial detergent formulations with high-temperature optima have been produced from Humicola lanuginosa (Lipolase) [4]. As the quantity of this enzyme was less than required for its commercial application, therefore, the yield of enzyme was increased by cloning the gene coding for this lipase in Aspergillus oryzae. This fungus produced increased quantity of the enzyme that can be used commercially in detergents [40]. The first commercial lipase, Lipolase, was introduced by Novo Nordisk in 1994. This enzyme was produced from Trichoderma lanuginosus and was expressed in Aspergillusz oryzae [83]. Novozymes later introduced three variants of Lipolase: Lipolase® Ultra, LipoPrime™, and Lipex® [13]. A presoak formulation was developed using an alkaline lipase produced by Trichosporon asahii, used for removing oil stains at ambient temperature [84]. The application of lipase from Aspergillus niger as an additive in a laundry detergent was reported by Saisubramanian et al. [85]. The enzyme showed increased stability in the presence of SDS, Tween 80 and in all commercial detergents. The washing process was optimized by Response Surface Methodology and the optimized conditions were 1% of commercial detergent, 75U of lipase, pH 9.5 and a washing temperature of 25 ◦C. Under these conditions, 33% of olive oil was removed from cotton fabric [85]. Several fungi like Aspergillus oryzae, Candida sp., Rhizopus oryzae, and Humicola lanuginosa are known to produce lipases under standardized conditions suitable for detergent applications [36].

3. Agrochemicals Fine and intermediate chemicals makers emphasize new products and processes for the pesticide industry via lipases, in view of its potential for decreasing costs and environmental contamination [86]. A variety of pesticides (insecticides, herbicides, fungicides or their precursors) having optically active compound are made using lipases [87-95]. Generally,

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these compounds were produced through the resolution of racemic mixtures of alcohol or carboxylic esters. Mitsuda and Nabeshima [87] have been used different fungal lipases for the transesterification of a racemate (R,S)-4-methyl-1-heptyn-4-en-3-ol, used as insecticide. Several fungal lipases were used for the lipase-catalyzed enantioselective hydrolysis of the acetic acid ester of racemic a-cyano-3- phenoxybenzyl alcohol (CPBA) for the production of (S)- CPBA, an active insecticidal stereoisomer [88]. The lipase-catalyzed resolution of 2-(4propan-2- yloxy)-phenyl propionic acid, an intermediate in the synthesis of a chiral acaricide, was described by Bosetti et al. [89], using lipase from yeast, fungus and bacteria. The stereoselective enantio-discrimination of Candida rugosa lipase yielded optically pure propionic acid derivative in the (S)-form. The (S)-form was then converted to the corresponding (R)-form, which was found to be very effective as an ovicide against the pest Tetranychus urticae [90]. The diastereomers of 4-hydroxyproline represent important building blocks for several agrochemicals and pharmaceuticals. Candida antarctica lipase B was identified among 43 different commercial lipases and esterases as an efficient biocatalyst for the enantioselective hydrolysis of racemic 4-oxo-1,2-pyrrolidinedicarboxylic acid dimethyl ester [91]. The final compound cis-4-hydroxy-D-proline or trans-4-hydroxy-Ddiastereomeric excess. Lipases from Candida cylindracea have been applied to the resolution of 2-bromopropionic acid and 2Chloropropionic acid which are starting materials for the synthesis of phenoxy propionic herbicides [43]. Optically pure amines can be used in the fine chemical industry as resolving agents, chiral auxiliaries, and chiral synthetic building blocks for pharmaceuticals as well as agrochemical compounds [92,93]. Wen et al. (2008) studied the enantioselective aminolysis of immobilized extracellular lipase from Yarrowia lipolytica by catalyzing enantioselective acylation of (±)-α-phenylethyl amine with acetic ester. In their optimized condition i.e. in hexane containing 3% DMSO at 45 °C, the enantiomeric excess of the product markedly increased from 0.35 to 0.96 after 6 days of reaction [94]. Shmidt et al. have reported the use of Candida antarctica lipase in reactions with stereospecific N-acylamines to prepare optically active amines [95].

4. Biopolymers Biopolymers like polyphenols, polysaccharides and polyesters show a considerable degree of diversity and complexity. Furthermore, these compounds are receiving increasing attention because they are biodegradable and produced from renewable natural resources. Lipases and esterases are used as biocatalysts for polymeric synthesis with the major advantages being their high selectivity (e.g. stereoselectivity, regioselectivity and chemoselectivity) under mild reaction conditions [96]. Lipase-catalyzed polymerization of polyesters is mainly carried out by two methods: Ring Opening Polymerization (ROP) and Polycondensation [97]. The key step of the ROP is the reaction of the lactone ring with lipase to provoke the ring opening and the formation of an acyl-enzyme intermediate. Nishida et al. Mentioned that Poly(1,4-dioxan-2-one) with an Mw of 41 400 was produced by an immobilized lipase from the Candida antarctica [98]. Berrera-Rivera et al. investigated the ring-opening polymerization reaction of the ε-caprolactone in the presence of

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n-heptane by the extracellular lipase from Yarrowia lipolytica. After 360 h of reaction at 50°C in the presence of 3mmol of ε-caprolactone and 100 mg of enzyme, a total conversion was obtained and polyester molecule presented an average mass of 970 Da [99]. It has been revealed that lipase can act as a powerful catalyst for the ROP of cyclic monomers and oligomers containing oxo-, thio-, and phosphoesters in addition to carbonate esters [97]. Feng et al. [100] reported that Candida rugosa lipase was successfully used in the enzymatic ring-opening copolymerization of ethyl ethylene phosphate and trimethylene Carbonate (TMC) to yield random copolymers having molecular weights ranging from 3200 to 10 200. The copolymerization of TMC with 15-pentadecanolide (PDL) was performed by Candida antarctica lipase in toluene at 70°C to yield random copolymers [101]. The enzymatic polymerization of ε-caprolactone with hydroxyethyl cellulose film for the production of hydroxyethyl cellulose-graft-poly(ε-caprolactone) was carried out by Candida antarctica lipase B [102,103]. Enantioselectivity for racemic lactones was carried out using lipase-catalyzed ROP of lactones to produce optically active polymers and optically active lactones that remained as unreacted material [97]. Lipases are also widely used for the enzymatic polymerization via polycondensation method [104]. First, enzymatic polymerization of dicarboxylic acids or their esters with glycols is described. For example, Mucor miehei lipase in immobilized form induced the polycondensation of adipic acid and 1,4-butanediol in diisopropyl ether [104]. The polymerization of isophthalic acid and 1,6-hexanediol using an immobilized lipase derived from Candida antarctica at 70 °C to produce an aromatic polyester was reported by Wu et al. [105]. Lipase from Candida antarctica induced the polymerization of divinyl esters of isophthalic acid, terephthalic acid, and p-phenylene diacetic acid with glycols to give polyesters containing aromatic moiety in the main chain [106]. Many authors mentioned the use of fungal lipases for polycarbonates synthesis via polycondensation. Diethyl carbonate was polymerized with 1,3-propanediol or 1,4-butanediol by Candida antarctica lipase catalysis in a successive two-step polymerization [107,108]. Functional polyesters were synthesized through the specific catalysis of lipase, and their properties and functions were evaluated. Enantio- and regioselective polycondensations produced chiral and sugar-containing polyesters, respectively [109,110]. An optically active oligoester was enantioselectively prepared from racemic 10-hydroxyundecanoic acid using lipase from Candida rugosa as catalyst. The resulting oligomer was enriched in the (S) enantiomer to a level of 60% enantiomeric excess (ee), and the residual monomer was recovered with a 33% ee favoring the antipode [111].

5. Biodiesel Production There are several reasons for the introduction of biodiesel as an alternative to conventional fossil based diesel. These include decreasing dependency on foreign energy supply from declining fossil fuel resources; helping to reduce global warming by using renewable biofuels for the transport sector; and lowering emissions of particles, sulphur, carbon monoxide and hydrocarbons [112,113].

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Biodiesel is produced by esterification of fatty acids or transesterification of oils and fats with short chain alcohols. Methanol is mostly used because of its lower cost compared with other alcohols. Several groups have reported the production of biodiesel through lipase catalysis [114,115]. De Oliveira et al. stated the use of two commercial fungal lipases, Novozym 435 and Lipozyme IM, as catalysts for the preparation of fatty acid ethyl esters from castor oil in hexane [116]. Novozyme 435 has also been used to catalyse the transesterification of crude soybean oils for biodiesel production in a solvent-free system [117]. Simple alkyl ester derivatives of restaurant grease were prepared using immobilized lipases from Thermomyces lanuginosa and Candida antarctica, as biocatalysts [118]. Pizarro and Park applied Rhizopus oryzae lipase powder (F-AP15) for methanolysis of vegetable oils from waste bleaching earths obtained from a crude vegetable oil refining process. The vegetable oils were extracted and identified as soybean, palm and rapeseed oil. Optimum conditions for methanolysis of extracted oils were 75% water content (by weight of substrate), an oil/methanol molar ratio of 1:4 and 67 IU/g of substrate with shaking of 175 rpm for 96 h at 35°C and the highest conversion yield reached 55% (w/w) was with palm oil after 96 h of reaction time [119]. Brusamarelo et al. have investigated soybean biodiesel production using the commercial product Novozym 435 within the temperature range of 45–70 °C, observing the highest yield (92%) when 65°C has been used [120]. Soetaert and Vandamme [121] reported the use of the lipases from Mucor miehei and Candida antarctica in the transesterification of various oils using hexane as solvent and found that the lipase from Mucor miehei is more efficient in converting primary alcohols (methanol, ethanol, propanol, and 1-butanol) with yields between 95% and 98%, whereas lipase from Candida antarctica is more proper for the conversion of secondary alcohols (isopropanol and 2-butanol) with yields between 61% and 84%. Ha et al. investigated the production of biodiesel in ionic liquids through immobilized Candida antarctica lipase-catalyzed methanolysis of soybean oil. The production yield reached 80% after 12 h at 50°C [122]. An immobilized lipase from Penicillium expansum was shown to be an efficient biocatalyst for biodiesel production from waste oil with high acid value in organic solvent. The production yield was 92.8% after 7 h of reaction [123]. The enzymatic production of biodiesel fuel from plant oils using Rhizopus oryzae cells immobilized within biomass support particles (BSP) for the methanolysis of soybean oil was investigated by Ban et al. [124]. When methanolysis was carried out with stepwise additions of methanol using BSP-immobilized cells, in the presence of 15% water the methyl esters content in the reaction mixture reached 90%-the same level as that using the extracellular lipase [124]. Adachi et al. mentioned that an Aspergillus oryzae whole-cell biocatalyst which coexpresses Fusarium heterosporum lipase (FHL) and mono- and di-acylglycerol lipase B (mdlB) in the same cell has been developed to improve biodiesel production. The reaction system using the lipase-coexpressing whole-cells was found to be superior in biodiesel production to others such as lipase-mixing and two-step reactions, affording the highest reaction rate (98%) [125].

6. Environmental Management Nowadays, industrial effluent treatment and environmental protection are two of the main industrial concerns, and must be dealt with in the most appropriate and cost-effective way, to

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avoid potential risks and costs. Wastewaters from dairies and slaughterhouses present high levels of fats and proteins, which can cause gross pollution of land and water [79]. The application of enzymes to transform or degrade specific pollutants is one of the more novel and innovative techniques in environmental management. Among these enzymes, lipases occupy a very important place in environmental applications due to their versatility and potentialities. The broadening use of lipases in bioremediation has achieved more importance to the removal of biofilm deposits from cooling water systems, to the manufacture of liquid soap, to the upgrading of waste fat, to bleaching and to the purification of waste gases expelled from factories [2]. The fascinating role of Aspergillus oryzae lipase in the degradation of waste hair for the production of cystine was another milestone in the bioremedial application of lipase [2]. In addition, Lipases are used in activated sludge and other aerobic waste processes, where thin layers of fats must be continuously removed from the surface of aerated tanks to permit oxygen transport. This skimmed fat-rich liquid is digested with lipases such as that from Candida rugosa [126]. Candida rugosa lipases were also used in the treatment of domestic wastewaters and in the cleaning of sewer systems, cesspools and sinkholes [69]. The Japanese company Meito Sangyo Co. produces a lipase from Candida rugosa (Lipase-MY) for fat removal in equipment of effluent treatment plants in the United States [127]. Jeganathan et al. [128] evaluate the hydrolysis of wastewater with high oil and grease concentration from a pet food industry using immobilized Candida rugosa lipase as a pretreatment step for anaerobic treatment through batch and continuous-flow experiments. Rigo et al. [129] stated the use of two fungal lipases, Penicillium restrictum lipase and Lipolase, in the hydrolysis of oil and grease in wastewater from the swine meat industry. Several studies mentioned the use of lipases for the degradation of wastewater contaminants from olive oil processing. De Felice et al. found that the yeast Yarrowia lipolytica was capable of reducing the COD value by 80% in 24 h and producing a useful biomass of 22.45 g/L as single cell protein and enzyme lipases [130]. Lipases have also been used to accelerate the biodegradation of polymers [131,132] and of slurries from oil-well perforations containing synthetic esters emulsified in water [133]. Poly(butylene adipate) (PBA) is a typical biodegradable synthetic plastic prepared by the polycondensation of 1,4-butanediol and adipic acid. PBA is readily transformed into the repolymerizable cyclic oligomers mainly consisting of BA dimer by lipase Candida antarctica [97]. Commercially available PBA with an Mw of 22 000 was degraded into a cyclic BA oligomer with an Mw of 600 by lipase Candida antarctica within 1 h. Poly(lactic acid) may be one of the most promising green plastics. Poly (L-lactic acid) (PLLA) having an Mw of 120 000 was transformed into cyclic oligomers by Candida antarctica lipase at 100 °C [97].

7. Biomedical Applications Interesterification and transesterification have great significance in pharmaceutical for selective acylation and deacylation [86]. In addition, chirality is a key feature in the efficiency of many pharmaceutical products, and consequently the production of single enantiomers of chiral intermediates has become increasingly significant in the pharmaceutical industry

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[134,135]. The ability of lipases to resolve racemic mixtures by the synthesis of a single enantiomer is currently exploited for drug production by the pharmaceutical industry. Profens, a class of nonsteroidal anti-inflammatory drugs, are active in the (s)-enantiomer form. Xie et al. synthesized pure (s)-ibuprofen using Candida rugosa lipase-catalyzed kinetic resolution via esterification [136]. Contesini et al. reported the successful use of Aspergillus niger lipase to resolve (R,S)-ibuprofen by a preferential esterification of (R)-ibuprofen in an organic solvent or in an ionic liquid [79]. Lipase from Candida antarctica (Novozyme (R) 435) has been used for the kinetic resolution of racemic flurbiprofen by the method of enantioselective esterification with alcohols [137]. Arroyo and Sinisterra reported that esterification reaction in non aqueous media using lipase-B from Candida antarctica was stereoselective towards the R-isomer of ketoprofen in a chiral solvent such as isobutyl methyl ketone and carvone [36]. Immobilized Candida antarctica lipase was also used as biocatalyst in the acylation reaction of the purine to synthesize a more soluble product having an antileukemia effect [138]. Hirashima et al. [36] have carried out the hydrolysis of acyl bonds at the 1-position of 1, 2 diacylglycerophospholipids for the purification of plasmalogens with Rhizopus delemar lipase. Candida rugosa lipase was used as biocatalyst in the esterification reaction of sulcatol and free fatty acid [139]. This biocatalyst was also used in the enzymatic resolution of some antimicrobial compounds ((S)- and (R)-elvirol and their derivates [140,141]. Palomo et al. [142] have immobilized Rhizopus oryzae lipase on dextran-hosphol coated Sepa beads and applied them to the enzymatic resolution of (±)-glycidyl butyrate which achieved an enantiomeric excess of >99 at 55% conversion. Enantiomers of alcohols, amines and carboxylic acids are very useful compounds in the pharmaceutical industry. Carvalho et al. [143] studied the kinetic resolution of (R,S)-2octanol with octanoic acid in n-hexane using four Aspergillus sp. lipase (A. flavus, A. niger, A. oryzae and A. terreus). They observed that lipases from Aspergillus niger and Aspergillus terreus are more enantioselective. Pilissão et al. [144] studied the enzymatic acylation of (R,S)- phenylethylamine, catalyzed by different lipases. It was observed that when Aspergillus niger lipase was used, in n-heptane at 35°C, a high conversion degree (30%) was obtained in 96 h of reaction with ethyl acetate as the acyl donor. Miyazawa et al. [145] reported the resolution of (R,S)-2-phenoxypropanoic acid using Aspergillus niger lipase by transesterification of the vinyl esters. The kinetic resolution of alpha-lipoic acid, which The (R)-enantiomer is much more active than the (S)-enantiomer against diabetes mellitus, HIV and tumors [79], was carried out using Aspergillus oryzae lipase [146].

8. Cosmetics The overwhelming interest of industrialists in screening lipases for use in the cosmetic and perfume industry has mainly been due to its activity in surfactants and in aroma production, the main ingredients in cosmetics and perfumes [2]. Immobilized Rhizomucor meihei lipase was used by Unichem International (Spain) as the biocatalyst for the production of isopropyl myristate, isopropyl palmitate and 2-ethylhexylpalmitate. The company claims that the use of the enzymatic process gives products of much higher quality, requiring minimum downstream refining [35].

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Monoacylglycerols and diacylglycerols, obtained by lipase esterification of glycerol, are useful as surfactants in cosmetics; they are added as texturing agents for improving the consistency of creamsand lotions [59]. Vitamin A and its derivatives (retinoids) have been used extensively in pharmaceuticals and cosmetics such as skin care products [147,148]. Immobilised Candida antarctica lipase (Novozym 435) was successfully used by Rajasse et al. for the acylation of retinol by reverse hydrolysis, alcoholysis, acidolysis and inter-esterification with several bifunctional acylating agents [149]. Retinyl palmitate was also synthesized in organic solvents with immobilized Candida sp. lipase [150]. Kojic acid, 5-hydroxy-2-(hydroxymethyl)-1, 4-pyrone, works widely in cosmetic preparations as a skin-lightening or bleaching agent [151]. However, it is hydrophilic and unstable in cosmetic use. In order to improve its lipophilicity, Liu et al. reported the synthesis of kojic acid esters via lipase-catalyzed esterification [152]. Lipase from Penicillium camembertii proved to be the best catalyst for the production of kojic acid monooleate. Under optimal conditions, the yield of kojic acid monooleate reached about 40%. After reaction at 40°C for 10 days, the lipase retained 57% of its initial activity [152].

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9. Biosensors Sensing lipids and lipid-binding proteins are a developing technology. In the fat and oil industry, in food technology and in clinical diagnosis, the quantitative determination of triacylglycerols is of great importance. Chemical methods for the analysis are rather costly and time-consuming. A promising new method involves the manipulation of microbial lipases as a biosensor. Lipases are fast, efficient, and accurate as sensors for the quantitative determination of triglycerides. The basic concept of using lipase as biosensors for lipids is to generate glycerol from triglycerides in the analytical sample and to quantify the released glycerol by enzymatic or chemical methods [35]. By screening different hydrolytic enzymes to fit the special demands, fungal lipases turned out to be the most practical [153]. Many authors have reported the use of a Candida rugosa lipase biosensor as a DNA probe [154,155]. Wei et al. [156] developed a method for the enzymatic determination of organophosphorous pesticides with a surface-acoustic-wave impedance sensor, which was based on lipase-catalysed hydrolysis. It was successfully applied to the determination of dichlorovos residues in the root, stem and blade of Chinese cabbage [157]. A potentiometric biosensor based on Candida rugosa lipase was developed by Kartal et al. [158]. This biosensor was applied for the detection of both triglycerides and organophosphorous pesticides [158].

10. Pulp and Paper Industry The use of enzymes in the pulp and paper industry has grown rapidly since the mid 1980s. Although many of these applications are still at the research and development stage, some of them have found their way into the mills in an unprecedented short period of time. One of the best examples is the enzymatic control of pitch in softwood mechanical pulps

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using fungal lipases, which was put into practice in a large-scale paper-making process as a routine operation in the early 1990s [159]. ‗Pitch‘ is a term used to describe collectively the hydrophobic components of wood, namely triglycerides and waxes, which cause severe problems in pulp and paper manufacture [160]. Nippon Paper Industries in Japan developed a pitch-control method that uses a fungal lipase from Candida rugosa to hydrolyse up to 90% of the triglycerides [161]. Lipases from Candida cylindracea have been shown to be effective in hydrolyzing triglycerides in extractives of fresh birch and birch sulfate pulp [162]. The enzyme treatment studies were continued by additional mill trials using an improved lipase preparation commercialized by Novo Nordisk (currently Novozymes, Bagsvaaerd, Denmark) under the trade name Resinase® [163-165]; a recombinant lipase expressed in Aspergillus oryzae. Resinase® hydrolyzed about 95% of the triglycerides in a pine mechanical pulp. In addition, the Resinase® treatment reduced the number of deposits, decreased the number of spots and holes in the paper, enabled a reduction in talc dosage to control pitch deposition and permitted the use of higher amounts of fresh wood. In addition to Resinase®, other industrial lipases – such as Lipidase 10000 (American Lab. Inc.) and Candida and Aspergillus lipases have been investigated for the enzymatic control of pitch [166].

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11. Textile Industry The use of fungal lipase in textile industry is becoming increasingly important. Lipases are used to assist in the removal of size lubricants in order to provide the fabric better absorbency for enhanced levelness in dyeing. It also reduces the frequency of cracks and streaks in the denim abrasion systems [36]. Polyethylene terephthalate (PET) is the most important synthetic fiber in the field. However, the PET fiber also has some undesirable properties due to its hydrophobic nature and its non-active surface. The treatment of PET fibers with lipases shows several advantages as compared to chemical methods. They act under mild and environmentally friendly conditions and do not harm the mechanical properties of the PET. Thus according to Wan et al. [167], Aspergillus oryzae lipase was capable of modifying PET fabrics, improving their hydrophilicity and anti-static ability. Several other studies have described the use and the effects of fungal lipases on different types of fabrics. Kim and Song demonstrated that the moisture regain of PET fabrics was considerably improved by fungal lipase treatment [168]. The enzymatic treatment of polyster fabric with Penicillium roqueforti lipase (Lipomod 338P) causes adequate effects, especially, referring to water penetration, absorption and the mechanical parameters of the processed fabric (strength, elongation, wear resistance) [169]. Kalantzi et al. reported that the use of Thermomyces lanuginosus lipase (lipolase) as a scouring agent for cotton fabrics enabled the creation of wax-free textiles where considerable amounts of pectin and protein were removed [170].

12. Leather Industry One of the oldest applications of industrial enzymes is processing hides and skins for leather. The various important processing methods involved in the leather manufacture are

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curing, soaking, liming, dehairing, bating, pickling, degreasing and tanning. All these successive steps in the leather production involve enzymatic action directly or indirectly for facilitating the procedures and enhancing the leather output of desired quality [171]. The application of lipases in different stages of the leather process, especially in degreasing, was previously studied [171]. Aspergillus lipase was previously used as biocatalyst in degreasing skins [172]. Yeshoda et al. [173] have used a fungal lipase for the degreasing of wooly sheep skins at 37°C under acidic conditions. Authors have also reported that degreasing and bating could be carried out simultaneously in the pH range of 7.8 to 8 [174]. It‘s also noteworthy that an acid lipase from Rhizopus nodosus was very effective in the degreasing of sheep skins [175].

REFERENCES [1] [2] [3] [4] [5] [6]

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[7] [8] [9] [10]

[11]

[12]

[13]

Kirk, O., Borchert, T. V., Fuglsang, C. C. Industrial enzyme applications. Curr. Opin. Biotechnol., 2002 13, 345-351. Pandey, A., Benjamin, S., Soccol, C. R., Nigam, P., Krieger, N. Soccol, V. The realm of microbial lipases in biotechnology. Biotechnol. Appl. Biochem., 1999 29, 119-131. Osborn, H. T., Akoh, C. C. Structured lipids, Novel fats with medical, neutraceutical, and food applications. Comp. Rev. Food Sci. Food Safety, 2002 3, 93-103. Jaeger, K. E., Ransac, S., Dijkstra, B. W., Colson, C., Heuvel, M., Misset, O. Bacterial lipases. FEMS Microbiol. Rev., 1994 15, 29-63. Schmid, R. D., Verger, R. Lipases: Interfacial Enzymes with Attractive Applications. Angew. Chem. Int., 1998 37, 1608-1633. Macrae, A. R., Hammond, R. C. Present and future applications of lipases. Biotechnol. Genet. Eng. Rev., 1985 3, 193-217. Davranov, K. Microbial lipases in biotechnology. Prikl. Biokhim. Mikrobiol., 1994 30, 527-534. Sharma, R., Chisti, Y., Banerjee, U. C. Production, purification, characterization and applications of lipases. Biotechnol. Adv., 2001 19, 627-662. Wiseman, A. Handbook of enzyme biotechnology. 3rd ed. Cornwall: Padstow; 1995. Dutra, J. C. V., Terzi, S. C., Bevilaqua, J. V., Damaso, M. C. T., Couri, S., Langone, M. A. P. Lipase production in solid state fermentation monitoring biomass growth of Aspergillus niger using digital image processing. Appl. Biochem. and Biotechnol., 2008 147, 63-75. Griebeler, N., Polloni, A. E., Remonatto, D., Arbter, F., Vardanega, R., Cechet, J. L., Isolation and screening of lipase-producing fungi with hydrolytic activity. Food Bioprocess Technol., 2009 4, 578-586. Grbavcic, S. Z., Dimitrijevic-Brankovic, S. I., Bezbradica, D. I., Siler-Marinkovic, S. S., Knezevic, Z. D. Effect of fermentation conditions on lipase production by Candida utilis. J. Serb. Chem. Soc., 2007 72, 757-765. Houde, A., Kademi, A., Leblanc, D. Lipases and Their Industrial Applications: An Overview. Appl. Biochem. Biotechnol., 2004 118, 157-170.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

128

Selmene Ouertani and Habib Horchani

[14] Tan, T., Zhang, M., Wang, B., Ying, C., Deng, L. Screening of high lipase producing Candida sp. and production of lipase by fermentation. Process Biochem., 2003 39, 459465. [15] Ellaiah, P., Prabhakar, T., Ramakrishna, B., Taleb, A. T., Adinarayana, K. Production of lipase by immobilized cells of Aspergillus niger. Process Biochem., 2004 39, 525528. [16] Larios, A., García, H. S., Oliart, R. M., Valerio-Alfaro, G. Synthesis of flavour and fragrance esters using Candida antactica lipase. Appl. Microbiol. Biotechnol., 2004 65, 373-376. [17] Cihangir, N., Sarikaya, E. Investigation of lipase producing by a new isolate of Aspergillus sp. World J. Microbiol. Biotechnol., 2004 20, 193-197. [18] Arai, T., Yusa, S., Kirimura, K., Usami, S. Cloning and sequencing of the cDNA encoding lipase I from Trichosporon fermentans WU-C12. FEMS Microbiol. Lett., 1997 152, 183-188. [19] Ota, Y., Gomi, K., Kato, S., Sugiura, T., Minoda, Y. Purification and some properties of cell bound lipase from Saccharomycopsis lipolytica. Agric. Biol. Chem., 1982 46, 2885-2893. [20] Mhetras, N. C., Bastawde, K. B., Gokhale, D. V. Purification and characterization of acidic lipase from Aspergillus niger NCIM 1207. Bioresource Technol., 2009 100, 1486-1490. [21] Saxena, R. K., Davidson, W. S., Sheoran, A., Giri, B. Purification and characterization of an alkaline thermostable lipase from Aspergillus carneus. Process Biochem., 2003 39, 239-247. [22] Dharmsthiti, S., Ammaranond, P. Purification and characterization of lipase from a raw-milk yeast (Trichosporon asteroids). Biotechnol. Applied Biochem., 1997 26, 111116. [23] Kakugawa, K., Shobayashi, M., Suzuki O., Miyakawa, T. Purification and characterization of a lipase from the glycolipid-producing yeast Kurtzmanomyces sp. I11. Biosci. Biotechnol. Biochem., 2002 66, 978-985. [24] Vakhlu, J., Kour, A. Yeast lipases: Enzyme purification, biochemical properties and gene cloning. Electron. J. Biotechnol., 2006 9, 69-85. [25] Diaz, M. J. C., Rodríguez, J. A., Roussos, S., Cordova, J., Abousalham, A., Carriere, F., Baratti, J. Lipase from the thermotolerant fungus Rhizopus homothallicus is more thermostable when produced using solid state fermentation than liquid fermentation procedures. Enz. Microbial Technol., 2006 39, 1042-1050. [26] Tan, T., Zhang, M., Xu, J., Zhang, J. Optimization of culture conditions and properties of lipase from Penicillium camembertii Thom PG-3. Process Biochem., 2004 39, 14951502. [27] Ben Salah, A., Sayari, A., Verger, R., Gargouri, Y. Kinetic studies of Rhizopus oryzae lipase using monomolecular film technique. Biochimie, 2001 83, 463-469. [28] Ben salah, R., Mosbah, H., Fendri, A., Gargouri, A., Gargouri, Y., Mejdoub, H. Biochemical and molecular characterization of a lipase produced by Rhizopus oryzae. FEMS Microbiol. Lett., 2006 260, 241-248. [29] Lima, V. M. G., Krieger, N., Mitchell, D. A., Fontana, J. D. Activity and stability of a crude lipase from Penicillium aurantiogriseum in aqueous media and organic solvents. Biochem. Eng. J., 2004 18, 65-71.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

Fungal Lipase

129

[30] Liu, Z., Zhenming, C., Lin, W., Jing, L. Production, purification and characterization of an extracellular lipase from Aureobasidium pullulans HN2.3 with potential application for the hydrolysis of edible oils. Biochem. Eng. J., 2008 40, 445-451. [31] Shu, C., Chun-Jun, X., Gee-Chen, L. Purification and partial characterization of a lipase from Antrodia cinnamomea. Process Biochem., 2006 41, 734-738. [32] Van Heerden, E., Litthauer, D., Verger, R. Biochemical characterisation and kinetic properties of a purified lipase from Aspergillus niger in bulk phase and monomolecular films. Enz. Microbial Technol., 2002 30, 902–909. [33] Pokorny, D., Cimerman, A., Steiner, W. Aspergillus niger lipases: induction, isolation and characterization of two lipases from a MZKI Al 16 strain. J. Mol. Catal. B: Enz., 1997 2, 215-222. [34] Maeda, H., Yamagata, Y., Abe, K., Hasegawa, F., Machida, M., Ishioka, R., Gomi, K., Nakajima, T. Purification and characterization of a biodegradable plastic-degrading enzyme from Aspergillus oryzae. Appl. Microbiol. Biotechnol., 2005 67, 778-788. [35] Hasan, F., Shah, A. A., Hameed, A. Industrial applications of microbial lipases. Enz. Microbial Technol., 2006 39, 235-251. [36] Singh, A. K., Mukhopadhyay, M. Overview of Fungal Lipase: A Review. Appl. Biochem. Biotechnol., 2009 166, 486-520 [37] Bosley, J. Turning lipases into industrial biocatalysts. Biochem. Soc. Trans., 1997 25, 174-178. [38] Aravindan, R., Anbumathi, P., Viruthagir, T. Lipase applications in food industry. Ind. J. Biotechnol., 2007 6, 141-158. [39] Chang, C. S., Tsai, S. W., Kuo, J. Lipase-catalyzed dynamic resolution of naproxen 2,2,2-trifluoroethyl thioester by hydrolysis in isooctane. Biotechnol. Bioeng., 1999 64, 120-126. [40] Hasan, F., Shah, A. A., Javed, S., Hameed, A. Enzymes used in detergents: Lipases. Afr. J. Biotechnol., 2010 9, 4836-4844. [41] Saxena, R. K., Ghosh, P. K., Gupta, R., Sheba Dvidson, W., Bradoo, S., Gulati, R. Microbial lipases, potential biocatalysts for the future industry. Curr. Sci., 1999 77, 101-115. [42] Theil, F. Lipase-supported synthesis of biologically active compounds. Chem. Rev., 1995 95, 2203-2227. [43] Sharma, D., Sharma, B., Shukla, A. Biotechnological Approach of Microbial Lipase: A Review. Biotechnol., 2011 10, 23-40. [44] Ferrer, M., Plou, F. J., Fuentes, G., Angelesgutes, M., Andersen, L., Kirk, O., Christensen, M., Ballesteros, A. Effect of the immobilization method of lipase from Thermomyces lanuginosus on sucrose acylation. Biocat. Biotrans., 2002 20, 63-71. [45] Ferrer, M., Soliveri, J., Plou, F. J., Lopez-Corter, N., Reyes-Duarte, D., Christensen, M., Copa-Patino, J. L., Ballesteros, A. Synthesis of sugar esters in solvent mixtures by lipase from Thermomyces lanuginosus and Candida antarctica B and their antimicrobial properties. Enz. Microb. Technol., 2005 36, 391-398. [46] Macrae, A. Enzymatic interesterification, Science Lecture Paper Series.1st edition. Befford, UK: Society of Chemical Industry; 2000. [47] Rajendran, A., Palanisamy, A., Thangavelu, V. Lipase Catalyzed Ester Synthesis for Food Processing Industries. Braz. Arch.Biology Technol., 2009 52, 207-219.

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130

Selmene Ouertani and Habib Horchani

[48] Gupta, R., Rathi, P., Bradoo, S. Lipase mediated upgradation of dietary fats and oils. Crit. Rev. Food Sci. Nutr., 43 2003, 635-644. [49] Pogori, N., Xu, Y., Cheikhyoussef, A. Potential Aspects of Lipases Obtained from Rhizopus Fungi .Res. J. Microbiol., 2007 2, 101-116. [50] Safari, M., Kermasha, S. Interesterification of butter fat by commercial lipase in a cosurfactant-free microemulsion system. J. Am. Oil Chem. Soc., 1994 71, 969-973. [51] Komatsu, T., Nagayama, K., Imai, M. Interesterification activity of Rhizopus delemar lipase in-hospholipids microemulsions. Coll. Surf. B: Biointerfaces, 2004 38, 175-178. [52] Bhattacharyya, S., Bhattacharyya, D. K. and De, B. K. (1999), Modification of tallow fractions in the preparation of edible fat, Weeks publishing Co.(www.foodproductdesign.com). [53] Australia Newzealand Food Authority, Lipase from genetically modified Aspergillus oryzae, Final Assessment Report, Application A41, May 2002. [54] Sahin, N., Akoh, C. C., Karaali, A. Lipase catalyze acidolysis of tripalmitin with hazelnut oil fatty acids and stearic acid to produce human fat substitutes. J. Agri. Food Chem., 2005 53, 5779-5783. [55] Tsuzuki, W. Acidolysis between Triolein and short chain fatty acid by lipase in organic solvents. Biosci. Biotechnol. Biochem., 2005 69, 1256-1261. [56] Klemann, L. P., Aji, K., Chrysam, M., D‘Amelia, R. P., Henderson, J. M., Huang, A. S., Otterburn, M. S., Yarger, R. G., Boldt, G., Roden, A. Random nature of Triacylglycerols produced by the catalyzed interesterification of Short and long chain fatty acid triglycerides. J. Agric. Food Chem., 1994 42, 442-446. [57] Khare, S. K., Nakajima, M. Immobilization of Rhizopus japonicus on celite and its application for enrichment of docosahexaenoic acid in soybean oil. Food Chem., 2000 68, 153-157. [58] Hiol, A., Jonzo, M. D., Rugani, N., Druet, D., Sarda, L., Comeau, L. C. Purification and characterization of an extracellular lipase from a thermophilic Rhizopus oryzae strain isolated from palm fruit. Enz. Microb. Technol., 2000 26, 421-430. [59] Carvalho, P. O., Campos, P. R. B., Noffs, M. D. A., Fregolente, P. B. L., Fregolente, L. V. Enzymatic Hydrolysis of Salmon Oil by Native Lipases: Optimization of Process Parameter. J. Braz. Chem. Soc., 2009 20, 117–124. [60] A. Sugihara, Y. Shimada, N. Takada, T. Nagao, Y. Tominaga, lipase: purification, characterization and its use for decosahexanoic acid enrichment of tuna oil. J. Ferment. Bioeng., 1996 82, 498–501. [61] Talon, R., Chastagnac, C., Vergnais, L., Montel, M. C., Berdagué, J. L. Production of esters by staphylococci. Int. J. Food Microbiol., 1998 45, 143-150. [62] Jackson, M. A., King, J. W. Lipase-catalyzed glycerolysis of soybean oil in supercritical carbon dioxide. J. Am. Oil Chem. Soc., 1997 74, 103-106. [63] Sonntag, N. O. V. Glycerolysis of fats and methyl ester status, reviews and critique. J. Am. Oil Chem. Soc., 1982 59, 795-802. [64] Ghamgui, H., Miled, N., Rebaï, A., Karra-chaâbouni, M., Gargouri, Y. Production of mono-olein by immobilized Staphylococcus simulans lipase in a solvent-free system: Optimization by response surface methodology. Enz. Microb. Technol., 2006 39, 717723. [65] Guo, Z., Xu, X. New opportunity for enzymatic modification of fats and oils with industrial potentials, Org. Biomol. Chem., 2005 3, 2615-2619.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

Fungal Lipase

131

[66] Ghosh, P. K., Saxena, R. K., Gupta, R., Yadav, R. P., Davidson, S. Microbial lipases: Production and applications, Sci. Prog., 79 (1996) 119-157. [67] Seitz, E. W. Industrial application of microbial lipases: a review. J. Amer. Oil Chem. Soc., 1974 51, 12-16. [68] Makhzoum, A., Owusu-Apenten, R. K., Knapp, J. S. Purification and properties of lipase from Pseudomonas fluorescens strain 2D. Int. Dairy J., 1996 6, 459-472. [69] Jaeger, K. E., Reetz, M. T. Microbial lipases form versatile tools for biotechnology. Trends Biotechnol., 1998 16, 396-403. [70] Cavaille-Lefebvre, D., Combes, D. Lipase synthesis of short-chain flavor thioesters in solvent-free medium. Biocatalysis Biotransform., 1997 15, 265-279. [71] Iwai, M., Okumura, S., Tsujisaka, Y. Synthesis of terpene alcohol esters by Iipase. Agric. Biol. Chem., 1980 44, 2731-2732. [72] Xu, Y., Wang, D., Mu, X. Q., Zhao, G. A., Zhang, K. C. Biosynthesis of ethyl esters of short-chain fatty acids using whole-cell lipase from Rhizopus chinensis CCTCC M201021 in non-aqueous phase. J. Mol. Cat. B: Enz., 2002 18, 29-37. [73] Shieh, C. J., Chang, S. W. Optimized synthesis of lipase catalyzed hexyl acetate in nhexane by response surface methodology. J. Agric. Food Chem., 2001 49, 1203-1207. [74] Chang, S. W., Shaw, J. F., Shieh, C. J. Optimization of enzymatically prepared Hexyl butyrate by lipozyme IM-77. Enzymatic synthesis of Hexyl Butyrate. Food Technol. Biotech., 2003 41, 237-243. [75] Loboret, F., Perraud, R. Lipase-catalyzed production of short-chain acids terpenyl esters of interest to the food industry. Appl. Biochem. Biotechnol., 1999 82, 185-198. [76] Greenough, R. J., Perry, C. J., Stavnsbjerg, M. Safety evaluation of a lipase expressed in Aspergillus oryzae. Food Chem Toxicol, 1996 34, 161-166. [77] Kumar, C. G., Malik, R. K., Tiwari, M. P. Novel enzyme-based detergents: An Indian perspective. Curr. Sci., 1998 75, 1312-1318. [78] Vulfson, E. N. Industrial applications of lipases in Lipases. Ed. By Wooley, P. and Petersen, S. B. Cambridge, Great Britain: Cambridge University Press; 1994. [79] Contesini, F. J., Lopesa, D. B., Macedoa, G. A., Da Grac, M., Nascimentob, A., De Oliveira Carvalhoc, P. Aspergillus sp. lipase: Potential biocatalyst for industrial use. J. Mol. Catal. B: Enz., 2010 67, 163-171. [80] Joseph, B., Ramteke, P. W., Thomas, G. Cold active microbial lipases: Some hot issues and recent developments. Biotechnol. Adv., 2008 26, 457-470. [81] Godfrey, T., West, S. The application of enzymes in industry. In: Godfrey, T., Reichelt, J., editors. Industrial enzymology. 2nd (edn). NewYork: The Nature Press; 1996; p. 512. [82] Aaslyng, D., Gormsen, E., Malmos, H. Mechanistic studies of proteases and lipases for the detergent industry. J. Chem. Technol. Biotechnol., 1991 50, 321-330. [83] Hoq, M. M., Yamane, T., Shimizu, S., Funada, T., Ishida, S. Continuous hydrolysis of olive oil by lipase in microporous hydrophobic membrane bioreactor. J. Am. Oil Chem. Soc., 1985 62, 1016-1021. [84] Kumar, S. S., Kumar, L., Sahai, V., Gupta, R. A thiol-activated lipase from Trichosporon asahii MSR 54: detergent compatibility and presoak formulation for oil removal from soiled cloth at ambient temperature. J. Ind. Microbiol. Biotechnol., 2009 36, 427-432.

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Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

132

Selmene Ouertani and Habib Horchani

[85] Saisubramanian, N., Edwinoliver, N. G., Nandakumar, N., Kamini, N. R., Puvanakrishnan, R. Efficacy of lipase from Aspergillus niger as an additive in detergent formulations: a statistical approach. J. Ind. Microbiol. Biotechnol., 2006 33, 669-676. [86] Stinson, S. C. Chiral Drugs. Chem. Eng. News, 1995 9, 44-74. [87] Mitsuda, S., Nabeshima, S. Enzymatic optical resolution of asynthetic pyrethroid alcohol. Enantioselective transesterification by lipase in organic solvent. Recl. Trav. Chim. Pays-Bas, 1991 110, 151-154. [88] Mitsuda, S., Yamamoto, H., Umemura, T., Hirohara, H., Nabeshima, S. Biological preparation of an optically active alcohol. Part III. Enantioselective hydrolysis of alphacyano-3-phenoxybenzyl acetate with Arthrobacter lipase. Agric. Biol. Chem., 1990 54, 2907-2912. [89] Bosetti, A., Bianchi, D., La-Porta, P., Bettarini, F. and Massimini, S. Biotechnology 94 – Applied Biocatalysis, 1994 68–70 [90] Bosetti, A., Bianchi, D., La-Porta, P., Bettarini, F., Massimini, S. Chemoenzymic synthesis of optically pure 1,3,4-Oxadiazol-2(3H)-ones with acaricidal activity. J. Agric. Food Chem., 1994 42, 2596–2599 [91] Sigmund, A. E., Hong, W., Shapiro, R., DiCosimo, R. Chemoenzymatic synthesis of cis-4-hydroxy-D-proline. Adv. Synth. Catal., 2001 343, 587-590. [92] Breuer, M., Ditrich, K.,Habicher, T., Hauer, B., Kesseler, M., Stuurmer, R. et al. Industrial methods for the production of optically active intermediates. Angew. Chem. Int., 2004 43, 788-824. [93] O'Donnell, M. The preparation of optically active alpha-amino acids from the benzophenone imines of glicine derivatives. Aldrichim. Acta, 2001 34, 3-15. [94] Wen, S., Tan, T., Yu, M. Immobilized lipase YlLip2-catalyzed resolution of alphaphenylethyl amine in a medium with organic cosolvent. Process Biochem., 2008 43, 1259-1264. [95] Smidt, H., Fischer, A., Fischer, P., Schmid, R. D. Preparation of optically pure chiral amines by lipase-catalyzed enantioselective hydrolysis of N-acyl-amines. Biotechnol. Tech., 1996 10, 335-338. [96] Gross, R. A., Kalra, B., Kumar, A. Polyester and polycarbonate synthesis by in vitro enzyme catalysis. Appl. Microbiol. Biotechnol., 2001 55, 655-660. [97] Matsumura, S. Enzymatic Synthesis of Polyesters via Ring-Opening Polymerization. In: Kobayashi, S., Ritter, H., Kaplan, D., editors. Enzyme-Catalyzed Synthesis of Polymers. Berlin-Heidelberg Germany: Springer-Verlag; 2006; 95. [98] Nishida, H., Yamashita, M., Nagashima, M., Endo, T., Tokiwa, Y. Synthesis of MetalFree Poly(1,4-Dioxan-2-one) by Enzyme-Catalyzed Ring-Opening Polymerization. J. Polym. Sci. Part A Polym. Chem., 2000 38, 1560-1567. [99] Berrera-Rivera, K., Flores-Carreon, A., Martinez-Richa, A. Enzymatic ring opening polymerization of ε-caprolactone by a new lipase from Yarrowia lipolytica. J. Appl. Pol. Sci., 2008 109, 708-719. [100] Feng, J., Zhuo, R., He, F. Sci. China Ser. B Chem., 2003 46, 160 [101] Kumar, A., Garg, K., Gross, R. A. Lipase-Catalyzed Copolymerizations of Trimethylene Carbonate and w-Pentadecalactone. Macromol., 2001 34, 3527-3533. [102] Gustavsson, M. T., Persson, P. V., Iversen, T., Hult, K., Martinelle, M. Polyester coating of cellulose fiber surfaces catalyzed by a cellulose-binding module - Candida antarctica lipase B fusion protein. Biomacromol., 2004 5, 106-112.

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Fungal Lipase

133

[103] Li, J., Xie, W., Cheng, H. N., Nickol, R. G., Wang, P. G. Polycaprolactone modified hydroxyethylcellulose films prepared by lipase catalyzed ring-opening polymerization. Macromol. 1999 32, 2789-2792. [104] Uyama, Hirosh, Kobayashi, S. Enzymatic Synthesis of Polyesters via Polycondensation. In: Kobayashi, S., Ritter, H., Kaplan, D., editors. Enzyme-Catalyzed Synthesis of Polymers. Berlin- Heidelberg Germany: Springer-Verlag; 2006; 133. [105] Wu, X., Linko, Y. Y., Seppälä, J. Lipase biocatalysis in the production of aromatic polyesters. Ann. N.Y. Acad. Sci., 1998 864, 399-404. [106] Uyama, H., Yaguchi, S., Kobayashi, S. Enzymatic Polymerization: A New Method of Polymer Synthesis. Polym. J., 1999 31, 380-388. [107] Matsumura, S., Harai, S., Toshima, K. (1999) Proc. Jpn. Acad. 75B:117 [108] Matsumura, S., Harai, S., Toshima, K. (2000) Macromol. Chem. Phys. 201:1632 [109] Uyama, H., Kobayashi, S. Enzyme-catalyzed polymerization to functional polymers. J. Mol. Catal. B Enz. 2002 117, 19-20. [110] Kobayashi, S., Uyama, H. Bio- and Biorelated Macromolecules. ACS Symp. Ser., 2003 840, 128-140. [111] O‘Hagan, D., Parker, A. H. Enzyme mediated polyester synthesis with the lipase from Candida rugosa preparation of an enantiomerically enriched polymer from an A-B monomer Polym. Bull. 1998 41, 519-524. [112] Demirbas, A. Progress and recent trends in biofuels. Prog. Energy Combust. Sci., 2007 33, 1-18. [113] Meher, L. C., Sagar, D. V., Naik, S. N. Technical aspects of biodiesel production by transesterification: A review. Renew. Sustain. Energ. Rev., 2006 10, 248–268. [114] Bajaj, A., Lohan, P., Jha, P. N., Mehrotra, R. Biodiesel production through lipase catalyzed transesterification: An overview. J. Mol. Catal. B: Enz., 2010 62, 9-14. [115] Robles-Medina, A., González-Moreno, P. A., Esteban-Cerdán, L., Molina-Grima, E. Biocatalysis: Towards ever greener biodiesel production. Biotechnol. Adv., 2009 27, 398-408. [116] De Oliveira, D., Di Luccio, M., Faccio, C., Rosa, C. D., Bender, J. P., Lipke, N. et al. Optimization of enzymatic production of biodiesel from castor oil in organic solvent medium. Appl. Biochem. Biotechnol., 2004, 113-116. [117] Du, W., Xu, Y. Y, Zeng, J., Jiu, D. H., Novozym 435-catalised transesterification of crude soya bean oils for biodiesel production in a solvent-free medium. Biotechnol. Appl. Biochem., 2004 40, 187-190. [118] Hsu, A. F., Jones, K., Foglia, T. A., Marmer, W. N. Immobilized lipase catalyzed production of alkyl esters of restaurant grease as biodiesel. Biotechnol. Appl. Biochem., 2002 36, 181-6. [119] Pizarro, A. V. L., Park, E. Y. Lipase-catalyzed production of biodiesel fuel from vegetable oils contained in waste activated bleaching earth. Process Biochem., 2003 38, 1077-1082. [120] Brusamarelo, C. Z., Rosset, E., Cesaro, A. Kinetics of lipase-catalyzed synthesis of soybean fatty acid ethyl esters in pressurized propane. J. Biotechnol., 2010 147, 108115. [121] Soetaert, W., Vandamme, E. J. Biofuels. Hoboken, NJ: Wiley; 2009.

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134

Selmene Ouertani and Habib Horchani

[122] Ha, S. H., Lan, M. N., Lee, S. H., Hwang, S. M., Koo, Y. M. Lipase-catalyzed biodiesel production from soybean oil in ionic liquids. Enz. Microbial Technol., 2007 41, 480483. [123] Li, N. W., Zong, M. H., Wu, Hong. Highly efficient transformation of waste oil to biodiesel by immobilized lipase from Penicillium expansum. Process Biochem., 2009 44, 685-688. [124] Ban, K., Kaieda, M., Matsumoto, T., Kondo, A., Fukuda, H. Whole cell biocatalyst for biodiesel fuel production utilizing Rhizopus oryzae cells immobilized within biomass support particles. Biochem. Eng. J., 2001 8, 39-43. [125] Adachi, D., Hama, S., Numata, T., Nakashima, K., Ogino, C., Fukuda, H., Kondo, Akihiko. Development of an Aspergillus oryzae whole-cell biocatalyst coexpressing triglyceride and partial glyceride lipases for biodiesel production. Biores. Technol., 2011 102, 6723-6729. [126] Bailey, J. E., Ollis, D. F. Applied enzyme catalysis. 2nd ed. In: Biochemical Engineering fundamentals. New York, NY: McGraw-Hill; 1986; 157. [127] Cammarota, M. C., Freire, D. M. G. A review on hydrolytic enzymes in the treatment of wastewater with high oil and grease content. Biores. Technol., 2006 97, 2195-2210. [128] Jeganathan, J., Nakhla, G., Bassi, A. Hydrolytic pretreatment of oily wastewater by immobilized lipase. J. Haz. Materials, 2007 145, 127-135. [129] Rigo, E., Rigoni, R. E., Lodea, P., Lodea, P., Oliveira, D., Freire, D. M. G., Treichel, H., Di Luccio, M. Comparison of two lipases in the hydrolysis of oil and grease in wastewater of the swine meat industry. Eng. Chem. Res., 2008 47, 1760-1765. [130] De Felice, B., Pontecorvo, G., Carfagna, M. Degradation of wastewaters from olive oil mills by Yarrowia lipolytica ATCC 20255 and Pseudomonas putida. Acta Biotechnol., 2004 17, 231-239. [131] Marten, E., Muller, R. J., Deckwer, W. D. Studies on the enzymatic hydrolysis of polyesters. I. Low molecular mass model esters and aliphatic polyesters. Polym. Degrad. Stab., 2003 80, 485-501. [132] Sivalingam, G., Chattopadhyay, S., Madras, G. Solvent effects on lipase catalyzed biodegradation of poly (epsilon-caprolactone) in solution. Polym. Degrad. Stab., 2003 79, 413-418. [133] Aliphat, S., Perie, F., Zurdo, C., Martingnon, A. 1998. Process for enzyme pretreatment of drill cutting. Patent US 5725771, United States. [134] Okuma, K., Ono, A. M., Tsuchiya, S., Oba, M., Nishiyama, K., Kainosho, M., Terauchi, T. Assymetric synthesis of (2S,3R)- and (2S,3S)-[2-13C;3-2H] glutamic acid., 2009 50, 1482-1484. [135] Kazi, B., Kiss, L., Forró, E., Fülöp, F. Synthesis of orthogonally protected azepane βamino ester enantiomers. Tetrahedron Lett., 2010 51, 82-85. [136] Xie, Y. C., Liu, H. Z., Chen, J. Y. Candida rugosa lipase catalyzed esterification of racemic ibuprofen and chemical hydrolysis of S-ester formed. Biotechnol. Lett., 1988 20, 455-458. [137] Zhang, H. Y., Wang, X., Ching, C. B., Wu, J. C. Experimental optimization of enzymic kinetic resolution of racemic flurbiprofen. Biotechnol. Appl. Biochem., 2005 42, 67-71. [138] [Alloue, W. A. M., Aguedo, M., Destain, J., Ghalfi, H., Blecker, C., Wathelet, J. P., Thonart, P. Les lipases immobilisées et leurs applications. Biotechnol. Agron. Soc. Environ., 2008 12, 57-68.

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Fungal Lipase

135

Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

.

[139] Janssen, E. M., Sjurenes, J. B., Vakurov, A. V., Halling, P. J. Kinetics of lipasecatalyzed esterification in organic media: correct model and solvent effects on parameters. Enz. Microb. Technol., 1999 24, 463-470. [140] [Ono, M., Suzuki, K., Tanikawa, S., Akita, H. First synthesis of (+)- and (–)-elvirol based on an enzymatic function. Tetrahedron Asymmetry, 2001 12, 2597-2604. [141] Kato, K., Ono, M., Akita, H. New total synthesis of (±)-, (–)- and (+)- chuangxinmycin. Tetrahedron, 2001 57, 10055-10062. [142] Palomo, J. M., Segura, R. L., Fernandez-Lorente, G., Guis, J. M., Fernandez-Lafuente, R. Enzymatic resolution of glycidyl butyrate in aqueous media. Strong modulation of the properties of the lipase from Rhizopus oryzae via immobilization techniques. Tetrahedron Asymmetry, 2004 15, 1157-1161. [143] Carvalho, P. O., Contesini, F. J., Bizaco, R., Macedo, G. A. Kinetic Properties and Enantioselectivity of the lipases Produced by four Aspergillus Wilds Species. Food Biotechnol., 2005b 19, 183-192. [144] Pilissão, C., Carvalho, P. O., Nascimento, M. G. Enantioselective acylation of (RS)phenylethylamine catalysed by lipases. Process Biochem., 2009 44, 1352–1357. [145] Miyazawa, T., Kurita, S., Shimaoka, M., Ueji, S., Yamada, T. Resolution of racemic carboxylic acids via the lipase-catalyzed irreversible transesterification of vinyl esters. Chirality, 1999 11, 554–560. [146] Yan, H., Wang, Z., Chen, L. Kinetic resolution of α-lipoic acid via enzymatic differentiation of a remote stereocenter. J. Ind. Microbiol. Biotechnol., 2009 36, 643648. [147] Keller, K. L., Fenske, N. A. Uses of Vitamins A, C, and E and related compounds in dermatology: A review. J. Am. Acad. Dermatol., 1998 39, 611-625. [148] Kligman, M. D. Topical treatments for photo aged ski. Postgrad. Med., 1997 102, 115118. [149] Rejasse, B., Maugard, T., Legoy, M. D. Enzymatic procedures for the synthesis of water-soluble retinol derivatives in organic media. Enz. Microb. Technol., 2003 32, 312-320. [150] Chunhua, Y., Tao, L., Tianwei, T. Synthesis of Vitamin A Esters by Immobilized Candida sp. Lipase in Organic Media. Chinese J. Chem. Eng., 2006 14, 81-86. [151] Burdock, G. A., Soni, M. G., Carabin, I. G. Evaluation of health aspects of kojic acid in food. Regul. Toxicol. Pharmacol., 2001 33, 80-101. [152] Liu, K. J., Shaw, J. F. Lipase-catalyzed synthesis of kojic acid esters in organic solvents. J. Am. Oil Chem. Soc., 1998 75, 1507-1511. [153] Kynclova, E., Hartig, A., Schalkhammer, T. Oligonucleotide labeled lipase as a new sensitive hybridization probe and its use in bio-assays and biosensors. J. Mol. Recognit., 1995 8, 139-145. [154] Pittner, F., Kynclova, E., Schalkhammer, T., Ecker, B., Wakolbingeer, W. (1995) Pat. AT-9302071 [155] Benjamin, S., Pandey, A. Isolation and characterization of three distinct forms of lipases from Candida rugosa produced in solid state fermentation. Braz. Arch. Biol. Technol., 2001 44, 213-221. [156] Wei, W., Wang, R. H., Nie, L. H., Yao, S. Z. Instrum. Sci. Technol. 1997 25, 157-167 [157] Wei, W., Wang, R. H., Nie, L. H., Yao, S. Z. Rapid determination of dimethoate with a surface acoustic wave impedance sensor system. Anal. Lett. 1997 30, 2641-2653

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[158] Kartal, F., Kilinc, A., Timur, S. Lipase biosensor for tributyrin and pesticide detection. Int. J. Environ. Anal. Chem., 2007 87, 10-11. [159] Hata, K. et al. Mill-scale application of enzymatic pitch control during paper production. In: Viikari, L., Jeffries, T. W., eds. Enzymes for Pulp and Paper Processing. ACS; 1996; pp. 280-296. [160] Farrell, R. L., Hata, K., Wall, M. B. Solving pitch problems in pulp and paper processes by the use of enzymes or fungi. Adv. Biochem. Eng. Biotechnol., 1997 57, 197-212. [161] [Reetz, M. T., Jaeger, K. E. Overexpression, immobilization and biotechnological application of Pseudomonas lipase. Chem. Phys. Lipids, 1998 93, 3-14. [162] Mustranta, A., Fagernäs, L., Viikari, L. Effects of lipases on birch extractives. Tappi J., 1995 78, 140-146. [163] Matsukura, M., Fujita, Y., Sakaguchi, H. On the use of ResinaseTM A for pitch control. Novo Publ. A, 1990 6122, 1-7. [164] Fujita, Y. Enzymic pitch control in papermaking process. Kami. Pa. Gikyoshi 1991 45, 905-921. [165] Fujita, Y. Recent advances in enzymic pitch control. Tappi J., 1992 75, 117-122. [166] Gutiérrez, A., Del Río, J. C., Jesús Martínez, M., Martínez, A. T. The biotechnological control of pitch in paper pulp manufacturing. Trends Biotechnol., 2001 19, 340-348. [167] Wan X., Lu D., Jönsson, L. J., Hong, F. Eng. Life Sci. 2008 8, 268–276. [168] Kim, H. R., Song, W. S. Lipase Treatment of Polyester Fabrics. Fib. Polym., 2006 7, 339-343. [169] Kim, H. R., Song, W. S. Lipase treatment to improve hydrophilicity of polyester fabrics. Int. J. Clothing Sci.Technol., 2010 22, 25-34. [170] Kalantzi, S., Mamma, D., Kalogeris, E., Kekos, D. Improved Properties of Cotton Fabrics Treated with Lipase and its Combination with Pectinase. Fib.Textiles East. Eur., 2010 18, 86-92. [171] Choudhary, R. B., Jana, A. K., Jha, M. K. Enzyme technology applications in leather processing. Ind. J. Chem.Technol., 2004 11, 659-671. [172] Trabitzsch, H., J. Soc. Leather Technol. Chem., 1966, 50, 382–389. [173] Yeshodha, K., Dhar, S. C., Santappa, M. Studies on the degreasing of skins using a microbial lipase. Leather Sci., 1978 25, 77-86. [174] Yeshodha, K., Dhar, S. C. and Santappa, M. (1978). Leather Science, 25, 267–273 [175] Muthukumaran, N., Dhar, S. C. Comparative studies on the degreasing of skins using acid lipase and solvent with reference to the quality of finished leathers. Leather Sci., 1982 29, 417-424.

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In: Lipase Editors: Hamdi Sassi and Sofien Cannamela

ISBN 978-1-62081-366-9 © 2012 Nova Science Publishers, Inc.

Chapter V

SECRETED PHOSPHOLIPASE A2 INHIBITORS FROM CYNARA CARDUNCULUS L.AND ALOE VERA EXTRACTS AS POTENTIAL THERAPEUTIC DRUG FOR INFLAMMATORY DISEASES Sofiane Bezzine* and Youssef Gargouri Laboratoire de Biochimie et de Génie Enzymatique des Lipases, BP « W » ENIS route de soukra, Sfax -Tunisia

ABSTRACT Copyright © 2012. Nova Science Publishers, Incorporated. All rights reserved.

The aim of the present work is to evaluate the anti-inflammatory properties of Cynara cardunculus L. (Asteraceae) during its growth and Aloe vera leaf skin. Cynara cardunculus L. and Aloe vera leaf skin (AVLS) were extracted using various solvents. The anti-inflammatory activities of crude extracts were evaluated by measuring the inhibition potency of mammalian non pancreatic phospholipases A2 (hG-IIA). The methanol and acetone extracts of leaves of C. cardunculus L. harvested on February exhibit potent inhibition of hG-IIA (IC50 = 50 µg/ml and 70 µg/ml, respectively). However, the acetone extract of stems harvested on December inhibit the hG-IIA with a lower IC50 around 130 µg/ml. Fractionation on silica gel and hydrophobic gel of the methanol extract of leaves of C. cardunculus L. harvested on February increase the inhibitory effect and the IC50 riched 10 µg/ml. Meanwhile, the water extract of AVLS exhibits the highest inhibitory effect with an IC50 = 0.22 mg/ml and interestingly no effect was observed on the digestive phospholipase A2 (group IB) even at a concentration of 5 mg/ml. Antioxidant activities of AVLS were also analyzed and the most active extracts were observed when using chloroform ethanol (1/1) and ethyl acetate (IC50 = 0.274 and 0.326 mg/ml, respectively). Analysis of the total phenolic content reveals that the water extract of AVLS, with the best anti-PLA2 effect, was poor in phenolic molecules (2 mg GAE/g). A significant correlation was established between the total phenolic content of AVLS and the antioxidant capacity but not with the anti PLA2 activity. *

Correspondence address: Pr. Sofiane BEZZINE, Laboratoire de Biochimie et de Génie Enzymatique des Lipases, ENIS route de soukra, 3038 Sfax-Tunisia. E-mail: [email protected].

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INTRODUCTION Phospholipases A2 (PLA2; EC 3.1.1.4) hydrolyze stereo specifically the sn-2 bond of 1,2-diacyl sn-3-phosphoglycerides generating fatty acids and lysophospholipids. Most of the PLA2 have been identified based on their gene sequences. They have been classified mainly into three groups: (i) cytosolic PLA2 (cPLA2); (ii) Ca2+-independent intracellular PLA2 (iPLA2) and (iii) Ca2+-dependent secreted PLA2 (sPLA2). They show different substrate specificity, Ca2+ requirement and they are key players in phospholipid metabolism. (1, 2). Secreted PLA2s are small proteins (14-19 kDa) with a rigid tertiary structure having five to eight disulfide bonds, which probably confer resistance to proteolysis and thermal denaturation (1, 2), and with a highly conserved catalytic site containing an His–Asp dyad and a Ca2+ binding loop. They were first detected in pancreatic juice and venoms. Secreted PLA2s are also expressed in a number of cell types and present in various body fluids. They participate in phospholipid catabolism as well as in antimicrobial defence against bacteria and other pathogens. The extensive literature on sPLA2s in inflammatory diseases has been exhaustively reviewed (3-10). The gene of pancreatic PLA2-IB was isolated in 1986 (11), followed in 1989 by the cloning of non-pancreatic PLA2-IIA from rheumatoid arthritic synovial fluid (12) and blood platelets (13). The sPLA2-IB and the sPLA2-IIA are the best known and biochemically characterized PLA2. This is the reason why our review will focus on these two PLA2 groups. Novel sPLA2 were identified by screening gene sequence databases. Eleven sPLA2 groups have been identified in mammals (IB, IIA, IIC, IID, IIE, IIF, V, X, III, XIIA and XIIB). They display partial overlapping tissue distribution (14, 15). Only sPLA2-IB and group X have an N-terminal prepropeptide and its proteolytic cleavage is a regulatory mechanism to generate an active enzyme (16). sPLA2-IB was found in large amounts in the pancreas and its main function is the digestion of dietary lipids (16). However, members of the group IB of sPLA2 are also found in non digestive tissues, including lung, spleen, gonad and kidney (11, 15). sPLA2-IIA has been first localized in the intestinal mucosa (17, 18) and in the synovial fluid of patients with rheumatoid arthritis as well as in platelets (13, 19). The concentration levels of PLA2-IIA increase in sera of patients suffering from severe acute inflammatory diseases such as sepsis, bacterial infections (20, 21) and acute pancreatitis (22). The sPLA2IIA was originally localized in Paneth cells of the rat intestine (23, 24) and later on in macrophages (25, 26). The two above mentioned cell types are both involved in the antibacterial response. Afterward it was demonstrated that the sPLA2-IIA from human and mouse, with high activity on phosphatidylglycerol and bearing cationic properties (pI > 9.0), are highly bactericidal against gram positive bacteria (27, 28) by perturbing the anionic bacterial cell wall (29). The bacteria killed by the intestinal PLA2 do not necessarily have to be within the intestinal lumen. Some bacteria specifically invade the intestinal mucosa from the lamina propria, as it has been postulated to occur in Whipple‘s disease (30). The human PLA2-IIA enzyme shows low affinity for zwitterionic lipids and in the absence of interfacial binding to membranes no hydrolysis is possible (31, 32). Based on the central role of lipid mediators in the inflammatory processes, the potential value of controlling phospholipid metabolism through PLA2 inhibition has always been acknowledged (33). Numerous compounds have been proposed as inhibitors of various

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sPLA2. However, clinical studies have never reached a therapeutical stage. The aim of this review was to evaluate some biochemical activities of extracts of Cynara cardunculus and Aloe vera such as their antioxidant capacity and their inhibitory effects on the proinflammatory phospholipase A2 group IIA. As a control experiment, we used the digestive pancreatic phospholipase A2 group IB. In fact, the plant Cynara cardunculus commonly named cardoon is known for its therapeutical uses in folklore as diuretic and choleretic activities (34, 35). Previous chemical investigations demonstrated the presence of saponins (36), sesquiterpene (37), lactones (38) and lignans (39) in this plant. Moreover, many aspartic proteases are also present in Cynara cardunculus. They are known as cardiosins, cyprosins, cenprosins and cynarases. The aim of this study was to evaluate the anti-inflammatory properties of different extracts of Cynara cardunculus L. during its growth by measuring the inhibitory effects on the hG-IIA phospholipase A2 by using various solvents for the extraction. Moreover, the Aloe vera L. (syn.: Aloe barbadensis Miller) is a perennial succulent plant belonging to the Aloeaceae family (sub-family of the Asphodelaceae) (40). Among over 400 Aloe species, Aloe vera is the most accepted specie for various medical, cosmetic and neutraceutical purposes (41-43). The plant is made of turgid green leaves joined at the stem in a rosette pattern. Each leaf consists of two parts: an outer green rind (skin) and an inner clear pulp (gel). The plant was described to contain a large amount of phenolic compounds (41, 4449) with a high content of 1,8-dihydroxyanthraquinone derivatives (aloe emodin) and glycosides (aloins), which were used as cathartic (50-52). Various studies have revealed that Aloe vera leaf skin (AVLS) possesses many pharmaceutical properties, including purgative (53), antibacterial (54, 55), anticancer (56-58), antifungal (59) and antioxidant (60-64).

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MATERIAL AND METHODS Enzymes and Reagents Human group IIA phospholipase A2 (hG-IIA) and Me-Indoxam were generous gifts from Dr. Gerard Lambeau (IPMC, France). Porcine group IB phospholipase A2 (pG-IB), Egg yolk phosphatidylcholine or lecithin, red phenol and sodium taurodeoxycholate (NaTDC), 2,2Diphenyl-1-picrylhydrasyl (DPPH), potassium phosphate, BHT (butylated hydroxytoluene), α-tocopherol and gallic acid were purchased from Sigma Chemical Co. (St.Loui, MO). Potassium ferricyanide, ferric chloride and Folin-Ciocalteu phenol reagent were purchased from Merck. Visible spectra measurements were performed using Anadéo visible spectrophotometer (Anadéo-Bibby).

Extractions The leaves and the stems of Cynara cardunculus L. were collected in the region of Sfax (South of Tunisia) on December, February and May. Dried and powdered leaves and stems were extracted successively using a soxhlet apparatus with n-hexane, dichloromethane,

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acetone and methanol. All extracts were concentrated to dryness in vaccuo and stored in the dark at 4 °C until use. Mature fresh leaves of Aloe vera (Aloaceae) with an approximate length of 0.5 to 0.7 m were collected from the region of Kairouan, (Tunisia) and harvested in August 2008. The fresh Aloe vera leaf skin (3 kg) was washed with distilled water and was extracted with 5 liters of ethanol (95 %) by maceration for 48 hours at room temperature. After a filtration step, the ethanolic extract was concentrated under reduced pressure and lyophilized to yield the ethanolic extract (79.80 g). This extract was suspended in water (200 ml) and partitioned successively with hexane (1 liter), ethyl acetate (4 liters), chloroform-ethanol (1/1, v/v) (1.5 liters) and butanol (1.5 liters). The remaining solution is designated ―water extract‖.

Fractionation of Compounds One gram of methanol extract of leaves of Cynara cardunculus was applied to a silica gel column. A mixture of chloroforme/methanol in a different volume ratio performed the elution (0/1; 1/4; 2/3; 3/2; 4/1; 1/0). Resulting fractions were labelled 1-6, evaporated and then tested for the PLA2 inhibitory effect. Fraction 2 (109 mg) was found to be the most active fraction. This fraction was applied to a medium pressure reverse phase column (C8) using acetonitrile/water (1/4; 1/1; 1/0). To achive the chromatography, the column was washed with methanol. Four fractions were collected, evaporated, labelled 2a-2d and again tested for their inhibitory effect on PLA2.

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Assay of sPLA2 Activity The sPLA2 (IIA and IB) activity were measured by colorimetry at 558 nm (65). Briefly, 3.5 mM of Egg yolk phospholipids were suspended in 0.1 M NaCl, 0.01 M CaCl2, 3 mM NaTDC (sodium taurodeoxycholate) and 0.055 mM red phenol; pH 7.6. The PLA2 activity was expressed in international units, one phospholipase unit corresponds to 1 µmol of titratable released fatty acid per minute under described conditions. For inhibition experiments, the sPLA2 (0.2 µg) was mixed and incubated at room temperature for 20 min with variable amounts of crude extracts before the measurement of PLA2 residual activity. Control experiments were performed with the corresponding amounts of solvent without extract. The residual activity of the sPLA2 and the IC50 values were then determined. The positive control measurement was carried out with Me-indoxam known as a potent covalent inhibitor of sPLA2.

Total Phenols Determination The total phenolic content (TPC) of the fractions of Aloe vera leaf skin (AVLS) was estimated by a colorimetric assay, according to the method described by Singleton and Rossi (1965) with some modifications (66). Briefly, 1 ml of sample at 1 mg/ml was mixed with 1 ml of Folin-Ciocalteu reagent. After 3 min of incubation, 1 ml of saturated Na2CO3 solution was added and the volume was adjusted to 10 ml with distilled water. The reaction mixture

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was kept in the dark for 90 min, after which the absorbance was read at 725 nm. The TPC was determined using gallic acid as a standard.

Assays of Antioxidant Activity The antioxidant activity was determined by using the following methods:

DPPH Radical Scavenging Assay The antioxidant activity was measured as equivalent of hydrogen-donating or radical scavenging ability, using the DPPH method (67-69) with some modifications. Briefly, 1.5 ml of DPPH solution at 10-5 M was incubated with 1.5 ml of extracts containing variable amounts of dry weight (between 0.01 and 1 mg). The reaction mixture was shaken and incubated in the dark for 30 min at room temperature. The control experiment was performed as described above without adding any extract. The absorbance (A) of the solution was measured at 517 nm. The radical scavenging activity was calculated using the following equation:

Scavenging effect (%) = (

1

ASamp le ACo n tro l

) x 100

(1)

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The extract concentration providing 50 % inhibition (IC50) was calculated from the plot of the scavenging effect against the extract concentration. BHT and α-tocopherol were used as standards.

Reducing Power Assay The reducing power of AVLS fractions was determined according to the method of Oyaizu (1986) (70). Solutions of variable concentration of AVLS extracts were mixed with 1 ml of 0.2 M sodium phosphate buffer at pH 6.6 and 1 ml of 1 % potassium ferricyanide (K3Fe(CN)6). The obtained reaction mixture was then incubated at 50 °C for 20 min. Next, 1 ml of 10 % (w/v) trichloroacetic acid was added to the mixture which centrifuged at 3000 rpm for 10 min. The upper layer solution (2.5 ml) was mixed with 2.5 ml of deionised water and 0.5 ml of fresh ferric chloride at 0.1%. The absorbance was measured at 700 nm. Higher absorbance of the reaction mixture indicates greater reducing power.

Statistical Analysis and Correlation Study Experimental results were given as mean value ± SD of three separate experiments. Statistical analysis was conducted using Microsoft Excel software. Differences at P < 0.05, using student‘s t-test, were considered to be significant. Correlation study was performed using the Graphic Pad Prism software, version 5.01.

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Sofiane Bezzine and Youssef Gargouri Table 1. Extraction yields of Cynara cardunculus L. with various solvent Organs

Month of Harvest

Hexane CH2Cl2 Acetone MeOH Hexane CH2Cl2 Acetone MeOH Hexane CH2Cl2 Acetone MeOH Hexane CH2Cl2 Acetone MeOH Hexane CH2Cl2 Acetone MeOH Hexane CH2Cl2 Acetone MeOH

December

Leaves

February

May

December

Stems

Solvent

February

May

Yield g/kg 23.0 36.0 24.2 57.9 14.8 26.8 15.5 36.0 27.7 44.4 31.5 57.7 9.7 9.8 10.3 44.2 7.6 7.5 4.4 59.4 22.4 9.1 5.0 142.0

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Table 2. Phytochemical screening and yields of AVLS extracts Fractions Yields (%) Steroids Terpenoids Carotenoids Anthraquinones Flavonoids Alkaloids Catechin tannins

Hexane 5.10 ++ ++ + -

Ethyl acetate 11.20 + ++ + ++ -

Chloroform-ethanol 6.09 +++ -

Butanol 8.57 + -

Water 66.67 +++

The yields were determined as relative to the initial amount of dry ethanolic extract (79.8 g). -: Absent, +: Present, ++: Rich, +++: Very rich.

RESULTS Extraction Yields of Cynara Cardunculus and Aloe Vera Leaves and stems of Cynara cardunculus were collected in different months (December, February and May). They were harvested and extracted by different solvents (n-Hexane, CH2Cl2, Acetone and Methanol) (Table 1). However, Aloe vera leaf skin was harvested in august. Table 2 show the various phytochemical families present in AVLS fractions. Tests for

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steroids, terpenoids and carotenoids were positive in both hexane and ethyl acetate fractions of AVLS. Anthraquinones were mostly detected in the ethyl acetate, chloroform-ethanol and butanol fractions while alkaloids and flavonoids were absent in all fractions. It is worth noticing that catechin tannins were only detected in the aqueous extract of AVLS.

Total Phenolic Contents of Aloe Vera Total phenolic contents from the various extracts of AVLS, expressed as milligram of GAEs per gram of extract, were presented in table 3. Among the five extraction systems used, the chloroform-ethanol extract showed the highest amount of phenolic compounds (40.5 mg GAE/g) and the poorest one was the water extract which contained only 2.07 mg GAE/g.

Antioxidant Activity of Aloe Vera

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DPPH Radical Scavenging Activity The antiradical activities of extracts were determined using the DPPH free radical assay. After the measurement of absorbance at 517 nm, the radical scavenging activities of the various extracts were expressed as the mean of the IC50 values (mg/ml). IC50 values of the five AVLS extracts, α-tocopherol and BHT are reported in table 3. Our results show that the chloroform-ethanol (1/1) extract of leaf skin exhibits the highest capacity to reduce DPPH (IC50 = 0.274 mg/ml), followed by the ethyl acetate extract (IC50 = 0.326 mg/ml) and the hexane extract (IC50 = 0.366 mg/ml). The lowest antiradical capacity was found in the water extract. For the sake of comparison, we measured the IC50 value of BHT and α-tocopherol which were 69 and 7.5 µg/ml respectively.

Table 3. Total phenol content and IC50 on DPPH of AVLS extracts Fractions

TPC (mg GAE /g of extract)

IC50 on DPPH radical (mg/ml)

Hexane

9.600 ± 0.014

0.366

Ethyl acetate

23.800 ± 0.058

0.326

Chloroform-ethanol (1/1) 40.500 ± 0.041

0.274

Butanol

16.900 ± 0.039

0.635

Water

2.072 ± 0.002

>1

BHT

-

69

-tocopherol

-

7.5

IC50 values on DPPH were calculated from the plot of the scavenging effect against the extract concentration. Effects of BHT and α-tocopherol, used as standard, were measured in the same conditions and expressed on µg/ml. All experiments were performed in triplicate ± standard deviation.

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Reducing Power Assay Results presented in Figure 1 show the reducing power of the AVLS fractions and BHT, used as reference. As expected, the reducing power of all samples tested is proportional to their concentrations. Below 0.2 mg/ml no significant difference was seen among all extracts. However differences were more pronounced with extracts at concentrations higher than 0.5 mg/ml. According to the results presented in Figure 1, ethyl acetate, chloroform-ethanol and butanol extracts were found to be better radical reducer (with electron donating capacities) as compared to water and hexane extracts. However, these extracts are less effective than BHT, a widely used commercial antioxidant which exhibits a high reducing capacity even when used at very low concentrations (0.05 mg/ml).

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Figure 1. Reducing power of AVLS measured with hexane, chloroform-ethanol, ethyl acetate, butanol and water extracts as compared to BHT. Experiments were performed in triplicate ± standard deviation.

A)

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B)

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Figure 2. Evaluation of the anti-PLA2 activity of leaves (A) and stems (B). Three harvests of air-leaves and stems of Cynara cardunculus.L were carried out on different months: The methanol extract of leaves harvested on February have an IC50 value of 0.05 mg/ml and the one of acetone extract is of 0.07 mg/ml. The acetone extract of stems harvested on December and February show a potent inhibition with an IC50 of 0.13 and 0.18 mg/ml, respectively. The reference molecule used on inhibitory test was the Me-Indoxam. In the same conditions, the IC50 of inhibition effect of the MeIndoxam on hG-IIA is 0.185 µg/ml (0.4 µM).

Evaluation of the PLA2 Inhibitory Effect Leaves of Cynara Cardunculus The inhibitory effect of leaves was evaluated using the method described in material and methods. Results reported in figure 2A show that the n-hexane and the dichloromethane extracts of leaves did not show any significant inhibitory effects on the hG-IIA phospholipase whatever big difference due to the month of harvest was observed. However, the acetone and the methanol extracts showed good inhibitory effects. The most potent inhibitions, corresponding to an IC50 value of 0.05 mg/ml and 0.07 mg/ml for the methanol and the acetone extracts, respectively; were observed when leaves were collected on February. Stems of Cynara Cardunculus L. The inhibitory effect of stems was evaluated using the same method A as previously described. Figure 2B shows that the extracts obtained using n-hexane and dichloromethane have no inhibitory effect on hG-IIA phospholipase. However, inhibition was observed in acetone and methanol extracts. In fact, the methanol extract inhibits the hG-IIA with an IC50 of 300 µg/ml when stems are harvested on February and 200 µg/ml when they are harvested on May.

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Meanwhile, Acetone extracts shows an IC50 value of 130 and 180 µg/ml when stems are collected on December and February, respectively. Methanol extract of leaves collected on February with IC50 of 0,05 mg/ml on hG-IIA was chosen as the best fraction and we evaluated their effect on the pancreatic sPLA2 (pGIB). Our results demonstrate that these extracts inhibit pG-IB with an IC 50 value of 500 µg/ml. We can conclude that inhibitory effect is more selective for hG-IIA more then pG-IB. We used the methanol extract of dry leaves harvested on February for fractionation. During purification, fractions were tested for the inhibitory effect on hG-IIA and pG-IB. Fraction 2 eluted from silica gel, which has an IC50 of 50 µg/ml, was subjected to a C8 chromatography. The elution from this hydrophobic column was performed using a mixture of acetonitrile/water. The most potent inhibitory effects were observed with fractions eluted at an acetonitrile /water: 1/1 and 1/0, respectively. The IC50 of these fractions are 10 and 20 µg/ml, respectively (data not shown).

Experiments were performed in triplicate ± standard deviation. Figure 3. IC50 of AVLS extracts measured during the inhibition of hG-IIA and pG-IB.

Table 4. Correlation significance between the total phenolic content of AVLS, antioxidant capacity and PLA2 inhibition Parameter

Number of XY Pairs Spearman r P value (one-tailed) P value summary Exact or approximate P value? Is the correlation significant? (alpha=0.1)

5 0,9 0,0417

IC50 value of scavenging activity 5 -0,9 0,0417

*

*

Exact Yes

Exact Yes

Reducing power

IC50 value of hG-IIA inhibition 5 0,0513 0,475 ns Exact No

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IC50 value of pG-IB inhibition 5 -0,7826 0,0667 *

Exact Yes

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Leaf Skin of Aloe Vera To evaluate the potential anti-inflammatory activity of AVLS, we tested the inhibitory effects of various extracts using two secreted phospholipases: hG-IIA involved in the inflammatory process and the pG-IB which hydrolyzes dietary phospholipids. Our main objective was to find an extract which was able to inhibit selectively the proinflammatory phospholipase A2 group IIA with no or minimal inhibitory effect on the digestive phospholipase A2 group. Out of the 5 extracts screened, three of them (ethyl acetate, chloroform-ethanol (1/1) and water) showed significant results (Figure 3). It is worth noticing that the water extract shows the most promising results in inhibiting the catalytic activity of the hG-IIA with an IC50 of 0.22 mg/ml. In sharp contrast, using the same extract even at concentrations higher than 5 mg/ml, no inhibition of the phospholipase A2 activity of pG-IB was noticed. These results indicate a selective inhibition of the water extract against these two sPLA2. Correlation Significance Study It was worth studying the potential correlations between the phenolic content of AVLS extracts with their antioxidant capacity and their inhibition of phospholipase A2 activity, since it was previously reported that phenolic compounds contribute directly to the antioxidant activity (71) and to the anti-inflammatory effects (72). A correlation analysis was performed between the total phenolic content, the antioxidant activity and the phospholipase A2 inhibition described in the present study. Results reported in table 4 show a linear regression and a significant relationship between the total phenolic content and free radical scavenging or reduction power (r = 0.9, P< 0.05). As expected, our results indicate that in the presence of high concentrations of the phenolic compounds, the antioxidant activity increases significantly. Furthermore, a positive correlation was also noticed between the total phenolic content and their inhibitory effect on pG-IB (r = 0.78, P< 0.1) but no significant correlation was obtained between the phenolic content and the hG-IIA inhibition.

DISCUSSION Anti-Inflammatory Activity of Cynara Cardunculus L To evaluate the anti-inflammatory effect of Cynara cardunculus L. on the biosynthesis of inflammatory mediators, different extracts were used to inhibit human proinflammatory PLA2. Results of this study showed that the percentage of inhibition depends on the nature of solvent used for extraction. In fact, we showed that inhibitory effect of extracts increases with the polarity of the solvent used. This is May be due to the presence of polar compounds such as lignans which are known for their anti-inflammatory activities (73). When polar solvents were used for the extraction such as acetone and methanol, differences on inhibitory effect were observed between leaves and stems. However, leaf extracts have more capacity to inhibit the hG-IIA phospholipase A2 than stems when using the methanol as extraction solvent whatever the month of harvesting. Using acetone for extraction, only the leaf extracts harvested on February shows a higher inhibitory effect than that of stems extract (IC50 of

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0.07 and 0.18 mg/ml, respectively). However, extracts of stems obtained on December (IC50 of 0.13 mg/ml) and May (IC50 of 1.6 mg/ml) show a better potenty to inhibit the sPLA2 than those of leaves (IC50 of 1.2 and 3.6 mg/ml respectively). In conclusion, methanol and acetone used for extraction of leaves harvested on February and the acetone extract of steams harvested on December showed a potent inhibition of the human sPLA2 involved in the inflammatory processes. Several studies investigated medicinal plants for potent natural inhibitors. From nine vine plants used in traditional Chinese medicine to treat inflammations, three of them exhibit a capacity to inhibit the PLA2 activity (74). The IC50 values obtained with ethanol extract of stem of Sinomenium acutum, Spatholobus suberectus and Trachelospermum jasminoide were 112, 54 and 33 µg/ml, respectively (74). Other study shows that the aqueous extract of Rhizophora mangle bark inhibited the sPLA2 with an IC50 value of 72µg/ml (75). Nunez et al., 2005 (76) have isolated the 4-nerolidylcatechol as an active principle that inhibit secreted phospholipase A2 group II. The enzyme activity of Bothrops asper myotoxin I was completely inhibited by 4-nerolylcatechol at an inhibitor: toxin ratio of 10:1 (wt/wt) with an IC50 of around 1mM. Compared to these works, our study on Cynara cardunculus extracts seems interesting. Purification and identification of sPLA2 inhibitors from Cynara cardunculus extracts are under investigation. Pharmacological study of the extract in animals is of high interest to develop new anti-inflammatory drugs.

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Anti-Inflammatory Activity of Aloe Vera The heterogeneity of the phytochemical composition of AVLS extracts is very promising for future medical applications. In fact, the chloroform-ethanol extract, which is rich in phenolic compounds (40.5 mg GAE/g), was the best antioxidant tested. In contrast, the water extract which is the poorest in phenolic compounds (2.07 mg GAE/g) did not show any antioxidant activity. This finding suggests that phenolic compounds from AVLS are responsible for the antioxidant effect. These results in agreement with those obtained by Kahkonen et al. (1999) (77), Shahidi and Marian (2003) (78) who reported that the differences in antioxidant activities of plant extracts could be due to the variable contents of their phenolic compounds. In the search for natural anti-inflammatory compounds, several studies were performed using Aloe vera due to its well known potent anti-inflammatory effects. Several authors have demonstrated the anti-inflammatory effect of this plant. Habeeb et al (2007) (79) have shown that the inner leaf gel from Aloe vera, when using an extract at a concentration of around 45 mg/ml, can suppress the cytokine induced inflammation after a whole bacterial stimulation of the human immune cells. We also report in this study that the water extract of AVLS possesses the best inhibitory effect on the pro-inflammatory PLA2 group IIA with an IC50 = 0.22 mg/ml. The phytochemical analysis showed that this water extract was very rich in catechin tannins suggesting that, apart from its low concentration in polyphenol contents, the molecules responsible for the anti-PLA2 effects belong probably to different chemical families. Probably the catechin tannins, rich in the water extracts and responsible for the anti-PLA2 activities, are different from the molecules bearing the antioxidant effect. It was previously reported that LY311727 inhibit hG-IIA and pG-IB with an IC50 of 0.47 and 8 µM,

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respectively (80). In the same case, Me-Indoxam was found to be the most potent inhibitor of sPLA2 and was extensively studied. It is able to inhibit hG-IIA with an IC50 of 0.006 µM and its IC50 on hG-IB is about 6µM (81) . Several studies evaluated the relationships between the antioxidant activity of plant extracts and their phenolic content. Velioglu et al. (1998) (82) reported a significant relationship between the total phenolic content and its antioxidant activity in selected fruits, vegetables and grain products. In agreement with these latter results, we report here a significant correlation between the total phenolic compounds and their antioxidant effects (r = 0.9, P