New Lipases And Proteases

NEW LIPASES AND PROTEASES

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NEW LIPASES AND PROTEASES

ABU BAKAR SALLEH RAJA NOOR ZALIHA RAJA ABDUL RAHMAN AND

MAHIRAN BASRI EDITORS

Nova Science Publishers, Inc. New York

Copyright © 2006 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. Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter cover herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal, medical or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Library of Congress Cataloging-in-Publication Data Abu Bakar Salleh. New lipases and proteases / Abu Bakar Salleh, Raja Noor Zaliha Raja Abdul Rahman ; and Mahiran Basri, editor. p. cm. Includes index. ISBN 978-1-60876-518-8 (E-Book) 1. Lipase. 2. Proteolytic enzymes. I. Noor Zaliha Raja Abdul Rahman, Raja. II. Mahiran Basri. III. Title. QP609.L5A28 2006. 572'.757--dc22 2006005503

Published by Nova Science Publishers, Inc. New York

CONTENTS Foreword

vii

Chapter 1

Lipases: Introduction Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri

1

Chapter 2

Protease: Introduction Abu Bakar Salleh, Che Nyonya Abdul Razak, Raja Noor Zaliha Raja Abd. Rahman and Mahiran Basri

23

Chapter 3

Thermostable Lipases Thean Chor Leow, Fairolniza Mohd Shariff, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri

41

Chapter 4

Organic Solvent Tolerant Lipases Syarul Nataqain Baharum, Mohamad Ropaning Sulong, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri

63

Chapter 5

Thermostable Proteases Noor Azlina Ibrahim, Thean Chor Leow, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri

77

Chapter 6

Organic Solvent Tolerant Proteases Azira Muhammad, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri

95

Chapter 7

Immobilized Enzymes Mohd Basyaruddin Abdul Rahman, Noor Mona Md. Yunus, Siti Salhah Othman, Abu Bakar Salleh and Mahiran Basri

111

Chapter 8

Modified Lipases Bimo Ario Tejo, Kok Whye Cheong, Abu Bakar Salleh and Mahiran Basri

127

List of Contributors

149

Index

151

FOREWORD It was in 1986 that a few of us got together to form the Enzyme & Microbial Research group at Universiti Putra Malaysia. We thought about palm oil which is a major commodity produced in Malaysia. As such it would be very critical to look at this vast resource and see how best it can be exploited. At the same time, biotechnology and environmental consciousness were becoming the issues of importance. So how could we add values to palm oil particularly with environmental friendly processes? Enzymatic processing of lipid and oils is becoming an important area of research. Hydrolytic enzymes, such as lipases and proteases are being sought after as the biocatalysts of the future. Synthetic reactions to create new compounds that have novel properties may be achieved through biocatalysis. Some of these compounds can replace or even improve on existing products that are produced by inorganic catalysis or being fractionated from unrenewable resources such as petroleum. We targeted on lipases and proteases as the most critical enzymes. Proteases are already widely used in industry and there is always scope for new proteases to be exploited in existing applications as well as new ones. Lipases have been projected as having exciting potential in the advancement of bioprocessing industry in particular oleochemicals. Thermostable enzymes are always being sought by the industries. Whereas solvent tolerant enzymes are becoming the vogue in view of their ability to function in low aqueous medium, suitable for synthetic reactions. This book presents work done in our laboratory on the search of new lipases and proteases. The work involves the broad range of biotechnology scope however, the focus of this book is on the search and acquisition, isolation and purification and the characterization of these enzymes. However, new enzymes include derivatised enzymes. Our laboratory is involved in the development and application of immobilized and modified enzymes. These designer enzymes may prove to be the new enzymes that would really expand the potential applications of enzymes in industries. We hope the information in this book enlightens our readers on our work and will be a useful reference in their work. We are most grateful to our students for their commitment and industriousness. We thank Universiti Putra Malaysia, the Ministry of Higher Education and the Ministry of Science, Technology and Innovation for their sustained support. Abu Bakar Salleh Raja Noor Zaliha Raja Abd.. Rahman Mahiran Basri Serdang, Malaysia

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 1-22

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 1

LIPASES: INTRODUCTION Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri ABSTRACT Lipases differ greatly with regard to their origins and their properties. Lipases can be obtained from microorganism, plant and animal. They can catalyze the hydrolysis or synthesis of a wide range of different substrates.. Microbial lipases are widely diversified in their enzymatic properties and substrate specificity which make them attractive for many industrial applications including detergents, food, flavor industry, ester and amino acid derivatives, baking, fine chemical, bioremediation, hard surface cleaning, leather and paper industry. Lipases stand amongst the most important biocatalysts due to their ability to utilize a wide spectrum of substrates, high stability towards extremes of temperature, pH and organic solvents, and chemo-, regio- and enantioselectivity. The enantioselective and regionselective nature of lipases have been utilized for the resolution of chiral drugs, fat modification, synthesis of cocoa butter substituents, biofuels, and for the synthesis of personal care products and flavor enhancers.

INTRODUCTION Lipases or triacylglycerol acylhydrolases (E.C. 3.1.1.3) are ubiquitous enzymes of considerable physiological significance and industrial potential that catalyze the hydrolysis of relatively long chain triacylglycerides (with acyl chain lengths of > 10 carbon atoms) with the formation of diacylglyceride, monoacylglyceride, glycerol and free fatty acids at the interface between the aqueous and organic phases (Thomson et al., 1999; Verger, 1997; Brockman et al., 1988). It is well known that the reaction is reversible and lipases can catalyze ester synthesis and transesterification in the reactions containing low water concentrations (Gupta et al. 2004; Saxena et al., 1999). Esters syntheses mediated by lipases have been under

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R. N. Z. R. A. Rahman, A. B. Salleh and M. Basri

scrutiny by numerous researchers in recent years as a wide variety of such compounds are very valuable. Compounds like trigylcerides, phospholipids, galactolipids, cutin, waxes, short chain esters and steroids play many diverse functions such as an energy sources, membrane constituents, emulsifiers, viscocity builders, protective coatings flavours. Functional properties of carboxylic esters are directly related to the length of the hydrocarbon backbone. Hence, esters of short and medium chain carboxylic acids and alcohol moieties play a relevant role in the food industry as flavor and aroma constituents. On the other hand, methyl and ethyl esters of long chain carboxylic acid moieties provide valuable oleochemical species that may function as fuel for the diesel engines, and esters of long chain carboxylic acid and alcohol moieties (typically referred to as waxes) have applications as lubricants and additives in cosmetic and pharmaceutical formulations. The hydrolytic and synthetic action of lipase is as shown Figure 1. O C

C O

C OH

(CH2)n CH3

C

C O

(CH2)n CH3

C

C

(CH2)n CH3

Triacylglycerol

+ 3 H2 O

Lipase

Water

O C

OH

C

OH

Glycerol

+

HO C

(CH2)n CH3

Fatty acid

Figure 1. Hydrolytic and Synthetic Action of lipase

The ability to catalyze hydrolysis of insoluble long chain fatty acid ester in the form of micelles, small aggregates or emulsion particles distinguishes lipases from other esterases which catalyze hydrolysis of soluble esters in preference to insoluble esters (Macrae, 1983). Apart from their natural substrates, lipases catalyze the enantio- and regioselective hydrolysis and synthesis of a broad range of natural and non-natural esters (Schmidt-Dannert, 1999). Lipases can attack the triacylglycerol molecule randomly, or may exhibit positional specificity (Marangoni and Rousseau, 1995). Nonspecific lipases that catalyze reactions at all positions in triacylglycerols are of limited value. On the other hand, the regiospecific lipases, e.g., the 1,3-specific lipases, offer the greatest potential in industrial applications, such as the production of structured lipids with unique functional properties (Sonnet and Gazzillo, 1991). Regiospecific lipases have great potential in the chemical, pharmaceutical, medical, cosmetics and leather industries (Iwai and Tsujisaka, 1984). There has been increasing interest in using enzymes, especially lipases, as hydrolytic or synthetic chiral catalysts (Hou and Johnston, 1992). Potential novel applications include the organic processing of optically pure compounds, use in digestive aids, and flavor and fragrance synthesis (Jarvis and Thiele, 1997). The enantioselectivity of lipases can also be exploited for the biosynthesis of a wide variety of chiral compounds (Qian and Xu, 2004; Kim et al., 2002; Shen et al., 2000). For example, Patel et al. (1996) used lipases to synthesize chiral drug intermediates for anticancer and other chemotherapeutic agents. Lipases have also proven useful for the modification or tailoring of fats. For example, the λ-linolenic acid content of borage oil can be enriched using a lipase-catalyzed reaction (Huang et al., 1997), while a cocoa butter equivalent can be

Lipases: Introduction

3

synthesized by lipase-catalyzed interesterification of the mid-fraction of palm oil (Mojovic et al., 1993). Lipases in the food-related industry have been used in the synthesis of amino acids derivatives. New and potential industrial applications such as these have increased the demand for new lipases, including those derived from microorganisms. Lipases are found in all living organisms, which constitute the two primary divisions of the phylogenetic tree - archaea (with a second division branching into the eukarya, including animals, plants and fungi) and bacteria (Olson et al., 1994). Lipolytic enzymes are indispensable for the biological breakdown of lipids. They are required as digestive enzymes in the transfer of lipids from one organism to another - from plant to animal and from animal to animal. Within the organisms, they are instrumental in the deposition and mobilization of fat as energy reserve, and in the metabolism of intracellular lipids and, therefore, in the functioning of the biological membranes. In eukaryotes, lipases are involved in various stages of lipid metabolism, including fat digestion, absorption, reconstitution and lipoprotein metabolism. Often the enzymes are found where their relevance is not very obvious, as in the venoms of reptiles and invertebrates and among extracellular microbial enzymes (Brockerhoff and Jenses, 1974).

SOURCE OF LIPASES Since lipases are physiologically necessary for living organisms, they are ubiquitous and can be found in diverse sources, such as plants, animals and microorganisms. More abundantly, however, they are found in bacteria, fungi and yeasts (Haki and Rakshit, 2003). Lipolytic enzymes are widely distributed in the plant kingdom, yet the knowledge of lipases from plants is still limited compared with those on mammalian and microorganism lipases (Vulfson, 1994). Many of the lipases so far studied in plants have been found to be membrane-bound and have been examined whilst still attached to the membranes. Lipases from maize, rape and castor bean have been solubilised and either purified to homogeneity or at least substantially purified (Vulfson, 1994). Three groups of lipolytic enzymes may be distinguished in mammals: the lipases discharged into the digestive tract by the specialized organs, the tissue lipases and the milk lipases (Gargouri et al., 1989). Several tissues and organs of mammals, such as the heart, brain, muscle, arteries, kidney, spleen, lung, liver, adipose tissue and serum, contain lipases (Vulfson, 1994). Microbial extracellular lipases are usually more thermostable than animal and plant lipases. In particular, lipases from thermophiles are expected to play a significant role in industrial processes since they are thermostable and resistant to chemical denaturation (Lee et al., 1999). In addition, the majority of lipases currently available is derived from mesophilic sources and display optimum activity at 35 to 40 °C (Sigurgísladóttir et al., 1993). Recently, microorganisms such as Geobacillus sp. T1 (Rahman et al., 2005; Leow et al., 2004), Bacillus sp. RSJ-1 (Sharma et al., 2001), Bacillus thermoleovorans ID-1 (Lee et al., 1999), Bacillus sp. THL027 (Dharmsthiti and Luchai, 1999), Bacillus thermocatenulatus (SchmidtDannert, 1994), Bacillus sp. strain A30-1 (Wang et al., 1995), Bacillus sp. Strain 398 (Kim et al., 1994), Bacillus spp. (Nawani et al., 1998; Becker et al., 1997; Llarch et al., 1997; Handelsman and Shoham, 1994; Sugihara et al., 1991), Pseudomonas cepacia (Sugihara et al., 1992), and Pseudomonas sp. (Kulkarni and Gadre, 1999), have reported as thermostable

4

R. N. Z. R. A. Rahman, A. B. Salleh and M. Basri

lipase producers. Besides, moderate thermophiles can be exploited to become good sources of thermostable enzyme as thermophiles (Fakhreddine et al., 1998). Knowledge of bacterial lipolytic enzyme is increasing at a rapid and exciting rate. Classification of bacterial esterases and lipases are based mainly on a comparison of the amino acid sequences and some fundamental biological properties. The grouping enable scientists to predict: (i) important structural features such as residues forming the catalytic site or the presence of disulphide bonds, (ii) types of secretion mechanism and requirement for lipase-specific foldase, and (iii) the potential relationship to other enzyme families (Jaeger and Eggert, 2002; Arpigny and Jaeger, 1999). According to Arpigny and Jaeger (1999), there are eight classes of bacterial lipolytic enzymes with the largest being further divided into seven subfamilies (Table 1): Lipases from subfamily I.1 of true lipases have molecular mass in the range of 30 – 32 kDa and display a higher sequence similarity to the P. aeruginosa lipases (Wohlfarth et al., 1991). In addition, lipases from subfamily I.2 are larger in size (33 kDa) owing to an insertion in the amino acid sequence which forms an anti-parallel double β-strand at the surface of the molecule. The lipases in subfamilies I.1 and I.2 depend on a chaperone protein named lipase specific foldase (‘Lif’) for folding. Lipase producers from subfamily I.3 secretes the enzyme with higher molecule mass than those from subfamilies I.1 and I.2 in one step through a three component ATP-binding-cassette transported system. In Bacillus lipases (Ruiz et al., 2002; Sinchaikul et al., 2001; Cho et al., 2000; Schmidt-Dannert et al., 1996), an alanine residue replaces the first glycine in the conserved pentapeptide: Ala – Xaa – Ser – Xaa – Gly. However, lipases from B. subtilis and B. pumilus stand apart because they are the smallest true lipases known (~20 kDa). Lipases from subfamily I.6 such as Staphylococcal haemolyticus lipase is a larger enzyme (~80 kDa) secreted as a precursor that is cleaved in the extracellular medium by a specific protease and processed rapidly into a 45 kDa mature lipase (Arpigny and Jaeger, 1999; Oh et al., 1999). Table 1: Families of lipolytic enzymes. Family

I

Subfamily

1

2

3 4

Enzyme-producing strain

Accession number

Pseudomonas aeruginosa Pseudomonas fluorescens C9 Vibrio cholerae Acinetobacter calcoaceticus Pseudomonas fragi Pseudomonas wisconsinensis Proteus vulgaris Burkholderia glumae Chromobacterium viscosum Burkholderia cepacia Pseudomonas luteola Pseudomonas fluorescens SIK W1 Serratia marcescens Bacillus subtilis Bacillus pumilus Bacillus licheniformis

D50587 AF031226 X16945 X80800 X14033 U88907 U33845 X70354 Q05489 M58494 AF050153 D11455 D13253 M74010 A34992 U35855

Similarity (%) Family Subfamily

100 95 57 43 40 39 38 35 35 33 33 14 15 16 13 13

Properties

True lipases

100 100 78 77 100 51 100 80 80

Lipases: Introduction

5

Table 1: Families of lipolytic enzymes (continued). Family

Subfamily

5

6

7 II (GDSL)

III

IV (HSL)

V

VI

VII

VIII

Enzyme-producing strain

Accession number

Geobacillus stearothermophilus Geobacillus thermocatenulatus Geobacillus thermoleovorans Staphylococcus aureus Staphylococcus haemolyticus Staphylococcus epidermidis Staphylococcus hyicus Staphylococcus xylosus Staphylococcus warneri Propionibacterium acnes Streptomuces cinnamoneus Aeromonas hydrophila Streptomyces scabies Pseudomonas aeruginosa Salmonella typhimurium Photorhabdus luminescens Streptomyces exfoliates Streptomyces albus Moraxella sp. Alicyclobacillus acidocaldarius Pseudomonas sp. B11-1 Archaeoglobus fulgidus Alcaligens eutrophus Escherichia coli Moraxella sp. Pseudomonas oleovorans Haemophilus influenzae Psychrobacter immobilis Moraxella sp. Sulfolobus acidocaldarius Acetobacter pasteurianus Synechocystis sp. Spirulina platensis Pseudomonas fluorescens Rickettsia prowazekii Chlamydia trachomatis Arthrobacter oxydans Bacillus subtilis Streptomyces coelicolor Arthrobacter globiformis Streptomyces chrysomallus Pseudomonas fluorescens SIK W1

U78785 X95309 AF134840 M12715 AF096928 AF090142 X02844 AF208229 AF208033 X99255 U80063 P10480 M57579 AF005091 AF047014 X66379 M86351 U03114 X53053 X62835 AF034088 AE000985 L36817 AE000153 X53868 M58445 U32704 X67712 X53869 AF071233 AB013096 D90904 S70419 S79600 Y11778 AE001287 Q01470 P37967 CAA22794 AAA99492 CAA78842 AAC60471

Similarity (%) Family Subfamily

15 14 14 14 15 13 15 14 12 14 14 100 36 35 28 28 100 82 33 100 54 48 40 36 25 100 41 34 34 32 20 100 50 24 20 16 100 48 45 100 43 40

100 94 92 100 45 44 36 36 36 100 50

Properties

Phospholipase

Secreted acyltransferase Secreted esterase OM-bound esterase OM-bound esterase Secreted esterase Extracellular lipase Extracellular lipase Extracellular esterase 1 Esterase Lipase Carboxylesterase Putative lipase Carboxylesterase Extracellular esterase 2 PHA-depolymerase Putative esterase Extracellular esterase Extracellular esterase 3 Esterase Esterase Carboxylesterase

Carbamate hydrolase p-Nitrobenzyl esterase Putative carboxylesterase Stereoselective esterase Cell-bound esterase Esterase III

The GDSL family does not exhibit the conventional pentapeptide Gly – Xaa – Ser – Xaa – Gly but rather display a Gly – Asp – Ser – (Leu) [GDSL] motif containing the active site serine residue. An extracellular lipase from Streptomyces rimosus R6-554W contained an ORF of 804 bp encoding a 268 amino acids polypeptide with 34 amino acid residues at the Nterminus, and the remaining polypeptide forming a mature lipase of 24.172 kDa (Vujaklija et

6

R. N. Z. R. A. Rahman, A. B. Salleh and M. Basri

al., 2002). Enzymes from family III display the canonical fold of α/β-hydrolases and contain a typical catalytic triad. The hormone-sensitive lipase (HSL) family displays a striking amino acid sequence similarity to the mammalian HSL. A cold-adapted lipase of an Alaskan psychrotroph, Pseudomonas sp. Strain B11-1 has consensus motifs conserved in mammalian hormone-sensitive lipase (Choo et al., 1998). Although the human-sensitive lipase (HSL) family and family V originate from mesophilic bacteria, cold adapted or heat adapted organisms, enzymes from family V share significant amino acid sequence similarity (20-25%) with various bacterial non-lipolytic enzymes, namely epoxide hydrolases, dehalogenases and haloperoxidases, which also possess the typical α/β-hydrolases fold and a catalytic triad. Enzymes from family VI are among the smallest esterases known with molecular mass around 23-26 kDa while rather large bacterial esterases (55 kDa) are normally found in family VII. Dimer formation is necessary to activate the enzyme from family VI. The subunit has the α/β-hydrolases fold and a classical Ser – Asp – His catalytic triad but the catalytic residues in the sequence (Ser-Asp-His) would not be expected in family VIII (Arpigny and Jaeger, 1999). In fact, lipases share very little homology especially upstream and downstream of open reading frame (ORF) except within the presumed catalytic region containing the conserved pentapeptide (Table 2). These conserved regions are essential for lipase catalysis (Dartois et al., 1992).

Table 2: Conserved catalytic sequence among microbial lipolytic enzymes. Organism Bacillus sphaericusa Bacillus subtilisb Bacillus stearothermophilusc Bacillus thermocatenulatusd Bacillus thermoleovoranse Staphylococcus haemolyticusf Staphylococcus warnerig Pseudomonas fragih Pseudomonas glumaei Pseudomonas aeruginosaj Pseudomonas fluorescensk Propionibacterium acnesl Moraxella sp.m

I V V V I I I V V V V V L

I D H H H H H N N N V D G

L I I I I L L L L L V F A

M V I I I I I I I I S V I

Protein sequence G E A H A H A H A H G H G H G H G H G H G H G H G W

S S S S S S S S S S S S S

A M Q Q Q M M Q Q H L Q M

G G G G G G G G G G G G G

G G G G G G G A G G G G G

Note: Residue which is strictly conserved in Bacillus spp. except Bacillus sphaericus 205y (in bold). References: a: Rahman et al., 2003 b: Dartois et al., 1992 c: Kim et al., 1998 d: Schmidt-Dannert et al., 1996 e: Cho et al., 2000 f: Oh et al., 1999 g: Kampen et al., 2001

h: Kugimiya et al., 1986 i: Frenken et al., 1992 j: Wohlfarth et al., 1992 k: Tan and Miller, 1992 l: Miskin et al., 1997 m: Feller et al., 1990

Lipases: Introduction

7

TYPES AND PROPERTIES Reactions of Lipases Lipases, (triacylglycerol acylhydrolases; EC 3.1.1.3) are one of the most important classes of hydrolytic enzymes that catalyze both the hydrolysis and the synthesis of esters (Sharma et al., 2001). They may or may not have positional specificity for the primary ester bonds. Hydrolysis of a triglycerids by lipases can yield di-and monoglycerides, glycerol and free fatty acids. In limited moisture, only esterification and transesterification are favored (Klibanov, 1997). Lipases also catalyze a number of useful reactions in a wide variety of organic solvents (Cernia and Palocci, 1997) such as esterification (Chowdary et al., 2001; Hamsaveni et al., 2001; Kiyota et al., 2001), transesterification, regioselective acylation of glycols and menthols, and synthesis of peptides (Zhang et al., 2001; Ducret et al., 1998) and other chemicals (Weber et al., 1999). Table 3 summarized various reactions catalysed by lipases. Table 3: Summary of various lipase reactions. HYDROLYSIS

O

O R1 C O

R2

+

R1 C OH

H2O

+

R2 OH

ESTER SYNTHESIS

O

O R1 C OH

+

R2 OH

R1 C O

R2

+

H2O

TRANSESTERIFICATION Alcoholysis

O

O R1 C O

R2

+

R3 CH2 OH

R1 C O

R3

R2 CH2 OH

+

Interesterification

O R1 C O

O

O R2

+

R3 C

O R4

R1 C O

O R4

R3 C

+

O R2

Acidolysis

O R1 C O

R2

+

O

O

R3 C OH

R3 C O

O R2

+

R1 C OH

The natural reaction of lipase as catalyst is the hydrolysis reaction on triglycerides. Hydrolysis process produces free fatty acid, soap and glycerol (Macrae and Hammond, 1985). Process catalyses by lipase occur in system containing emulsion of fatty acid in water or oil in water or soluble buffer where enzyme can be soluble. Fatty acid is widely used in industry but is found only in small quantities in the natural world. Studies on hydrolysis of olive oil

8

R. N. Z. R. A. Rahman, A. B. Salleh and M. Basri

(Shiomori et al., 1996), palm oil (Noor et al., 2003), milk fat (Patel et al., 1996), and short chain substrates (Nini et al., 2001) to produce fatty acids have been reported. Esterification is the reversible process of hydrolysis where equilibrium between forward and reverse reaction is controlled by water content in the reaction mixture. In low water environment or non-aqueous system, lipase acts efficiently to catalyse the reversible reactions including esterification where ester bonds formed between fatty acid and alcohol. In reaction system containing acid and alcohol, organic solvent may be added to dissolve reaction components in ester reaction where water is produced. To push the thermodynamic equilibrium to continuously produce ester, water must be removed. Through this process compounds such as ester terpene alcohol (Lee et al., 1998) and glyceride whether mono, di or triglyceride can be synthesized in a proportion that depends on that between the reactants and on the reaction conditions (Hilal et al., 2006; Blanco et al., 2004). Esterification by lipases appears to be an attractive alternative to bulk chemical routes. In fact, ester synthesis can be performed at room temperature and pressure, as well as neutral pH in reaction vessels operated either batch wise or continuously. Products obtained therefore are, qualitatively, more pure than the ones obtained by alternative chemical means because chemical catalysis tends to be unspecific and consequently generate several by-products. Therefore use of lipases to carry out esterification alleviates the need for a wide variety of complex postreaction separation processes (which are a must in chemical processes), and thus leads to lower overall operation costs. However, the drawbacks of the extensive use of lipases and biocatalysts in general compared to classical chemical catalysts can be found in the relatively low stability of enzyme in their native state as well as their prohibitive cost (Villeneuve et al., 2000). Interesterification process involves exchange of acyl radical between ester and acyl donor. If the acyl donor is free fatty acid, this reaction is called acidolysis. If it is between acid and alcohol, reaction is called alcoholysis, whereas reaction defined as interesterification is the process of modification the oil and fat composition and thus modifying the physical characteristics of the triglyceride mixture (Macrae, 1983). By interesterification reaction, certain fatty acid on a triglyceride is hydrolysed and replaced by another fatty acid by the esterification process (Macrae and Hammond, 1985).

Mechanism of Lipase-Catalyzed Reactions Though lipases differ widely in the number of amino acids in their primary sequences, yet the common feature of all lipases is that their active site is built up of the three amino acids serine, aspartate or glutamate and histidine (catalytic triad). The three-dimensional structure of all lipases follows a common motif, the α/β-fold of hydrolase, where lipase is consisting of eight mostly parallel β-sheets surrounded on both sides by α-helices (Kazlauskas, 1998) (Figure 2).

Lipases: Introduction

9

Figure 2. Schematic representation of α/β-fold of hydrolase

The triad and several oxyanion-stabilizing residues are thought to compose the active center of lipases. A helical segment (lid or flap), which blocks the active center, is responsible for the important characteristics of lipase-interfacial activation which let them to be distinguished from other hydrolases such as serine proteases or esterases. Based on the x-ray structure of lipase before and after binding to the substrate, researchers believe that lipases are activated before they take part in biochemical transformation reactions (Pleiss, 2000; Soumanou et al., 1999). A freely dissolved lipase in the absence of an aqueous/lipid interface resides in its inactive state and a part of the enzyme molecule covers the active site. However, when the enzyme contacts the interface of a biphasic water-oil system, a short α-helix (the lid) is folded back leading to activation of the lipase. In other words, with no substrate present, the lid is closed and the enzyme is inactive, whereas in the presence of substrate, the water-oil interface exists, and the lid is opened then lipase is active (Orru et al., 1999). The catalytic triad (Ser, His and Asp), the actual chemical operator in the active site, is shown in the mechanism of a serine-hydrolase-catalyzing hydrolysis of an ester (Figure 3) (Gotor, 2002; Marangoni, 1995; Estell 1993; Gotor, 1992; Gotor et al., 1991).

Substrate Specificity The hydrolysis of a substrate occurs at the oil-water interface. Thus, the velocity of the lipolysis of ester bonds is a function of the surface area of the substrate in the reaction mixture. However, the surface area differs with each substrate because of differences in the physio-chemical properties. Due to these factors, different sources of microbial lipases display different substrate specificity. Generally, lipases can be classified into three groups by their substrate specificity - positional, fatty acid and partial glycerides specificity.

a) Positional Specificity Hydrolysis of triglycerides releases diglycerides, monoglycerides, glycerol and free fatty acids. Lipases can be divided to two groups on the bases of their positional specificity. The first group is non-specific, releasing fatty acids from all three positions of the triglyceride randomly. The second group has 1, 3-positional specificity, releasing only the fatty acids from

10

R. N. Z. R. A. Rahman, A. B. Salleh and M. Basri

the outer 1- and 3-positions of the triglyceride. Only fatty acid in the outside position (at position 1 and 3) is cleaved by 1, 3 specificity lipase. Owing to steric hindrance, the middle fatty acid at position 2 is not reacted on. Positional specificity is retained if lipase is used in organic solvents. However, through random acyl migration, the 2-fatty acid monoglyceride undergoes rearrangement pushing the fatty acid to the 1 or 3 position of the glycerol molecule; as acyl migration is a slow process and as the available lipases do not act on glycerol 2-mono fatty acid esters, the hydrolysis slows down and awaits the acyl migration to complete for enabling the lipase to attack the glyceride at the 1 and/or the 3 position (Saxena et al., 1999).

Figure 3. The serine hydrolase mechanism; Step I. the special arrangement of these three groups performs a decrease of the pK-value of the serine hydroxy group thus enabling it to perform a nucleophilic attack on the carbonyl group of the substrate R1COOR2. Thus, the acyl moiety of the substrate is covalently linked onto the enzyme, forming the acyl enzyme intermediate and liberating the alcohol R2OH. Step II. water (regarded as the nucleophile, Nu.) attacks the acyl-enzyme intermediate, regenerating the enzyme and releasing the carboxylic acid R1COOH. Depending on the medium used (aqueous or organic medium) any other nucleophile can compete with water for the acyl-enzyme intermediate thus leading to a number of synthetically useful transformations. If the alcohol R4OH attacks the acyl-enzyme intermediate, the ester R1COOR4 is formed (interesterification reaction). But If the amine R3NH2 or hydrogen peroxide attack the acyl-enzyme intermediate, an amide R1CONHR3 (enzymatic aminolysis of esters) or the peracids R1COOH are formed respectively. Hydrazinolysis yields hydrazides and the action of hydroxylamine results in the formation of hydroxamic acid derivatives.

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Generally, the position specificity of a lipase is determined by using triolein as substrate in the reaction mixture. Hydrolysis of the substrate is carried out for a certain time, and then the products extracted with n-hexane and analyzed by thin layer chromatography. A silica gel plate developing in the mobile phase (solvent system) will show spots of the hydrolysis products either by exposure to iodine or by spraying with acid and heating. The lipases from Pseudomonas. aeruginosa LST-03 (Ogino et al., 2000) and BTID-B lipase from Bacillus thermoleovorans ID-1 (Lee et al., 2001) have been shown to split the ester bonds at all positions of the glycerol moiety, i.e. they are not position specific. Other bacterial lipases are 1, 3-position specific, producing end products of 2-monoolein and fatty acids. For example, the lipases from Bacillus sp. THL027 (Dharmsthiti et al., 1999), three strains of B. stearothermophilus, and three strains of B. licheniformis and one strain of B. subtilus (Chen et al., 2004) have been shown to be 1, 3-position specific.

b) Fatty Acid Specificity To determine the fatty acid specificity, monoacids and triglycerides are used. Sometimes using synthetic triglycerides is a problem since solid triglycerides can only be hydrolyzed slowly by microbial lipases (Sugiura and Isobe, 1974). The fatty acid specificities of microbial lipase from Streptomyces rimosus were studied using alkylnoate esters of varying alkyl chain lengths. There are some differences in the abilities of microbial lipases to hydrolyze fatty esters. For example, the highest hydrolysis rates were obtained by S. rimosus lipase with p-NPcaprylate (C8), p-NP-caprate (C10) and p-NP-laurate (C12), indicating the enzyme preference for medium acyl chain lengths. The p-NP esters of palmitic and myristic acids were also good substrates, but p-NP-stearate was only hydrolyzed at a considerably slower rate. Abramic et al. (1999) reported that short chain ester p-NP-butrate was a good substrate for purified enzyme. Sinchaikul et al. (2001) reported that Bacillus stearothermophilus P1 lipase hydrolyzed synthetic substrates with acyl group chain lengths between C8 and C12 with optimal activity on C10 p-NP-caprate. The lipase activity on long chain of substrates was between 70 and 100% optimal for C8 or C10 groups and 30 and 50 for C12 to C18, whereas, with short-chain substrates (C2-C6), the lipase activity was less than 30%. In addition, the lipase hydrolyzed triacylglycerols with acyl-group chain lengths between C8 and C12, with optimal activity on C8 (tricaprylin) and trilinolenin more than trilinolein and triolein. Kambourova et al. (2003) demonstrated that B. stearothermophilus MC 7 lipase has wide substrate specificity towards triglycerides with C4 to C18, the highest activity being on tributryrin. This enzyme hydrolyzed triglycerides with short-chain fatty acids more rapidly than those with long-chain fatty acids. However, the enzyme was also able to hydrolyze olive oil (which long-chain fatty acids). On the other hand, Pseudomonas. aeruginosa LST-03 lipase had the highest activity against tricaproin (C6) compared to the other triglycerides used (Ogino et al., 2000). Among various fatty acid methyl esters of different carbon chain lengths, the enzyme had the highest hydrolytic activity against methyl octanoate (C8). Furthermore, the lipase activity on long-chain natural oils were 55% and 60% those on olive oil (oleic acid) and castor oil (ricinolein acid), respectively, compared to a higher activity on coconut oil which principal fatty acid is lauric (C12). The lowest activity (12%) was obtained on tung oil (eleostearic acid).

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c) Partial Glycerides Specificity Another specificity of a lipase is its activity on the fatty acids in mono- and diglycerides. Sakiyama et al. (2001) reported that lipase from Pseudomonas sp. LP7315 showed hydrolytic activity against all the monoglycerides tested, but with the activity dependent on the type of monoglyceride. The highest activity was on monomyristin. The enzyme was not able to hydrolyze di- and triglycerides, including olive oil. This phenomenon indicates that Pseudomonas sp. LP7315 lipase had a strict specificity for monoglycerides. The lipase purified from Bacillus sp. H-257 showed the highest hydrolytic activity against monolaurin (Imamura and Kitaure, 2000).

APPLICATIONS OF LIPASES Lipases constitute an important group of biotechnologically valuable enzymes, mainly because of the versatility of their applied properties and ease of mass production (Hasan et al., 2006). Enzymes are of increasing commercial importance because of their broad substrate specificities, these valued biocatalysts act under mild conditions, highly stable in various organic solvents, reduced wastes produced in their use (Snellman et al., 2002). The outstanding characteristic of lipases is their affinity for the water-oil interface. The surface activity of the enzyme is an important determinant of its ability to catalyze reactions with hydrophobic substrates because it has to compete for the water-oil interface with a multitude of other proteins. When catalyzing the hydrolysis of triglycerides, lipases are known to be selective with regard to the chain lengths and double bond positions on the acyl groups, in addition to showing regio- and/or stereoselectivity in catalysis (Osterberg et al., 1989). Lipases, especially those of microbial origin, have great potential in commercial applications, such as in the production of food additives (for flavor modification), fine chemicals (synthesis of esters), detergents (hydrolysis of fats), waste water treatment chemicals (decomposition and removal of oil substances), cosmetics (removal of lipids), pharmaceuticals (digestion of oil and fats in foods), leather (removal of lipids from animal skins) and medical products (blood triglyceride assay) (Kamini et al., 2000). Lipases can be used to accelerate the degradation of fatty wastes (Masse et al., 2001) and polyurethane (Takamoto et al., 2001).

Detergents The major commercial application for hydrolytic lipases is their use in laundry detergents with nearly 32% of the total lipase sales. Thermostability and ability to remain active in the alkaline environment are necessary for lipases to be use in detergents formulation. Lipases find promising applications as additives in industrial laundry and household detergents due to their ability to hydrolyze fats. In detergent, lipases are especially selected to meet the following requirements: (1) a low substrate specificity, i.e., an ability to hydrolyze fats of various compositions; (2) ability to withstand relatively harsh washing conditions (pH 10–11, 30–60 °C); (3) ability to withstand damaging surfactants and enzymes, which are important ingredients of many detergent formulations. Novo Nordisk introduced the first commercial

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recombinant lipase ‘Lipolase,’ in 1994, which originated from the fungus Thermomyces lanuginosus and was expressed in Aspergillus oryzae. In the following year, two bacterial lipases were introduced—‘Lumafast’ from Pseudomonas mendocina and ‘Lipomax’ from P. alcaligenes—by Genencor International, AU-KBC Research Center, Life Sciences, Anna University, Chennai, India (Jaeger and Reetz, 1998). An alkaline lipase, produced by P. alcaligenes M-1, which was well suited to removing fatty stains under conditions of a modern machine wash was reported by Gerritse et al. (1998). A combination of continuous screening from various sources and protein engineering were shown to be valuable tool in obtaining lipases with desired properties (Cardenas et al., 2001; Kazlauskas and Bornscheuer, 1998; Wang et al., 1995; Yeoh et al., 1986).

Food Industry Lipases have unique characteristics, including substrate specificity, stereospecificity, regionselectivity and the ability to catalyze heterogeneous reactions at the interfaces of watersoluble and water-insoluble systems (Sharma et al., 2002). The esters produced play important roles in the food industry as flavor and aroma constituents (Pandey et al., 1999; Gandhi et al., 1995). The position of the fatty acid in the glycerol backbone, the chain length of the fatty acid, and its degree of unsaturation are important in determining the physical properties of a triglyceride with desired nutritional and sensory values. Upgrading of less desirable and relatively inexpensive fats to cocoa butter substitutes is conducted through lipase-catalysed transesterification reaction that replaces the palmitic acid in palm oil with stearic acid (Undurraga et al., 2001). In dairy and confectionery industries, lipases are used to hydrolyze milk fat to endow dairy products, particularly cheeses with desired specific flavors as a result of different carbon chain length free fatty acids. In addition, lipases are also used for development of particular flavors in bakery products, and beverages (Kazlauskas and Bornscheuer, 1998).

Pulp and Paper Industry Lipases are used together with cellulases and ligninases during paper manufacturing. Treatment of the pulp with lipase leads to a considerable improvement in productivity and a sustained high quality with less frequent clearing of the drying cylinders (BjÖrkling et al., 1991). Lipases are used to remove the pitch (hydrophobic components of wood) from the pulp during paper making. Nippon Paper Industries in Japan developed a pitch-control method that uses a fungal lipase from Candida rugosa to hydrolyze up to 90 % of the triglycerides (Sharma et al., 2001; Jaeger and Reetz, 1998).

Biodiesel Biodiesel (fatty acid methyl/ethyl esters), which is derived from triglycerides by transesterification with methanol/ethanol, has attracted considerable attention during the past decade as a renewable, biodegradable, and nontoxic fuel. Enzymatic transesterification using lipase has become more attractive for biodiesel fuel production, since the glycerol produced as a by-product can easily be recovered and the purification of fatty methyl esters is simple to accomplish. The production cost is significantly reduced by the use of whole cell biocatalysts

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immobilized within biomass support particles as no purification process is necessary (Hsu et al., 2002; Fukuda et al., 2001).

Crude Oil Refining Phospholipase A2 (secreted by the archaea Pyrococcus horikoshii) has been used in crude oil refining (Yan et al., 2000). The enzyme reacted optimally at 95oC and pH 7.0. Phospholipase A2 is a better degummer for oil refineries, reducing the wastewater problems and running cost (Klaus, 1998). Lipases from thermophiles are expected to play increasingly important roles in industrial processes as they are thermostable and resistant to chemical denaturation (Lee et al., 1999). Inherently stable biocatalysts are well appreciated and hence systematic screening on potential new organisms. Alternatively, molecular cloning incorporated with site-directed mutagenesis is being performed to produce recombinant enzymes with desired characteristics (Lllanes, 1999).

Other Applications Esters of long chain carboxylic acid and alcohol moieties (waxes), the products of lipase reactions have applications as lubricants and additives in cosmetics formulations (Linko et al., 1994). Lipases are endowed with a substrate specificity that surpasses any other known enzymes. Consequently, the industrial usage of lipases can be expected to increase rapidly in the future. Nowadays, lipases have been used to hydrolyze lipids to obtain fatty acids and glycerol for soap production. During processing of hides and skins, removal of residual fats and protein debris that are associated with the hide and the hair are being carried out by enzymatic processes instead of chemical process due to the latter’s inefficient liming and environmental problems. Besides, lipases are utilized in activated sludge and other aerobic waste process for removing thin layers of fats from the surface of aerated tanks to permit oxygen transport (Gandhi, 1997). Lipases are also being used to synthesize glucoside esters that are difficult to make using traditional chemical means. The compounds exhibit surfactant properties and may represent an attractive alternative to existing products in areas ranging from cosmetics, to household detergents, to industrial de-greasing of metals, electronics and leather. These “Green” chemicals are safer for the user, and are environmentally more acceptable due to a very rapid biodegradation and low toxicity (BjÖrkling et al., 1991). Lipases are also employed in the production of pharmaceuticals, cosmetics, agrochemicals and medical diagnostics (Gandhi, 1997). Lipase catalysed enantioselective reactions for the synthesis of pharmaceutical products as the chirality determines the inhibition effects of antitumour and antibacterial agents (Jaeger and Eggert, 2002). Besides, lipases are also used in the efficient production of enantiopure (S)-indanofan, a novel herbicide used against grass weeds in paddy fields (Tanaka et al., 2002). Other applications include the of non-cellulosic impurities from raw cotton before further processing with dyes into finished products and removal of subcutaneous fat in the leather industry (Pandey et al., 1999). The future demands for lipases would be greater as more products requiring the catalytic ability of lipases are being innovated. The ability for lipases to function in low water environment would further expand the potential for lipase applications. With the ability to discover and develop specifically designed enzymes, the use of lipases would surely be enhanced.

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Klaus, D., 1998. An enzyme process for the physical refining of seed oils. Chemical Engineering and Technology, 21: 278-281. Klibanov, A. M., 1997. Why are enzymes less active in organic solvents than in water? Trends in Biotechnology, 15: 97–101. Kugimiya, W., Otani, Y., Hashimoto, Y., and Takagi, Y., 1986. Molecular cloning and nucleotide sequence of the lipase gene from Pseudomonas fragi. Biochemical and Biophysical Research Communications, 141 (1): 185-190. Kulkarni, N., and Gadre, R. V., 1999. A novel alkaline, thermostable, protease-free lipase from Pseudomonas sp. Biotechnology Letters, 21: 897-899. Lee, D. W., Kim, H. W., Lee, K. W., Kim, B. C., Choe, E. A., Lee, H. S., Kim, D. S., and Pyun, Y. R., 2001. Purification and characterization of two distinct thermostable lipases from the gram-positive thermophilic bacterium Bacillus thermoleovorans ID-1. Enzyme and Microbial Technology, 29: 363-371. Lee, D. W., Koh, Y. S., Kim, K. J., Kim, B. C., Choi, H. J., Kim, D. S., Suhartono, M. T., and Pyun, Y. R., 1999. Isolation and characterization of a thermophilic lipae from Bacillus thermoleovorans ID-1. FEMS Microbiology Letters, 179: 393-400. Lee, K. K. B., Poppenborg, L. H., and Stuckey, D. C., 1998. Terpene ester production in a solvent phase using a reverse micelle-encapsulated lipase. Enzyme and Microbial Technology, 23: 253-260. Leow, T. C., Rahman, R. N. Z. A., Basri, M., and Salleh, A. B., 2004. High level expression of thermostable lipase from Geobacillus sp. Strain T1. Bioscience, Biotechnology, and Biochemistry, 68 (1): 96-103. Linko, Y., Lamsa, M., Huhatala, A., and Linko, P., 1994. Lipase catalyzed transesterification of rapeseed oil snf 2-ethyl-1-hexanol. Journal of the American Oil Chemists' Society, 71: 1411–1414. Llarch, A., Logan, N. A., Castellví, J., Prieto, M. J., and Guinea, J., 1997. Isolation and characterization of thermophilic Bacillus spp. from geothermal environments on deception island, south Shetlanh Archipelago. Microbiological Ecology, 34: 58-65. Lllanes, A., 1999. Stability of biocatalysts. Electronic Journal of Biotechnology, 2 (1): 1-9. Macrae, A. R., 1983. Extracellular microbial lipases. In: Microbial Enzymes and Biotechnology. (eds.) Fogarty, W. M., Applied Science Publishers Ltd. England, pp: 225250. Macrae, A. R., and Hammond, R. C., 1985. Present and future applications of lipases. Biotechnology and Genetic Engineering Reviews, 3: 193-219. Marangoni, A. G., and Rousseau, D., 1995. Engineering triacylglycerols: the role of transesterification. Trends in Food Science and Technology, 6: 329-335. Masse, L., Kennedy, K. J., and Chou, S. P., 2001. The effect of an enzymatic pretreatment on the hydrolysis and size reduction of fat particles in slaughterhouse wastewater. Journal of Chemical Technology and Biotechnology, 76: 629–35. Miskin, J. E., Farrell, A. M., Cunliffe, W. J., and Holland, K. T., 1997. Propionibacterium acnes, a resident of lipid-rich human skin, produces a 33 kDa extracellular lipase encoded by gehA. Microbiology, 143: 1745-1755. Mojovic, L., Siler, M. S., Kukic, G., and Vunjak, N. G., 1993. Rhizopus arrhizus lipase catalyzed interesterification of the midfraction of palm oil to a cocoa butter equivalent fat. Enzyme and Microbial Technology, 15: 438-443.

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Nawani, N., Dosanjh, N. S., and Kaur, J., 1998. A novel thermostable lipase from a thermophilic Bacillus sp.: characterization and esterification studies. Biotechnology Letters, 20 (10): 997-1000. Nini, L., Sarda, L., Comeau, L. C., Boitard, E., Dubès, J. P., and Chahinian, H., 2001. Lipasecatalysed hydrolysis of short-chain substrates in solution and in emulsion: a kinetic study. Biochimica et Biophysica Acta, 1534: 34-44. Noor, I. M., Hasan, M., and Ramachandran, K. B., 2003. Effect of operating variables on the hydrolysis rate of palm oil by lipase. Process Biochemistry, 39: 13-20. Ogino, H., Nakagawa, S., Shinya, K., Muto, T., Fujimura, N., Yasuda, M., and Ishikawa, H., 2000. Purification and characterization of organic solvent-stable lipase from organic solvent tolerant Pseudomonas aeruginosa LST-03. Journal of Bioscience and Bioengineering, 89 (5): 451-457. Oh, B. C., Kim, H. K., Lee, J. K., Kang, S. C., and Oh, T. K., 1999. Staphylococcus haemolyticus lipase: biochemical properties, substrate specificity and gene cloning. FEMS Microbiology Letters, 179: 385-392. Olsen, G. J., Woese, C. R., and Overbeek, R., 1994. The winds of (evolutionary) change: breathing new life into microbiology. Journal of Bacteriology, 176 (1): 1-6. Orru, R. V. A., Archelas, A., Furstoss, R., and Faber, K., 1999. Epoxide hydrolases and their synthetic applications. In: Biotransformations, Advances in Biochemical Engineering/Biotechnology. (eds.) Faber, K., Springer-Verlag. Heidelberg. Germany, 63: 145-167. Osterberg, E., Blomstrom, A. C., and Holmberg, K., 1989. Lipase catalysed transesterification of unsatorated lipids in amicroemultion. Journal of the American Oil Chemist’s Society, 66 (9): 1330-1333. Pandey, A., Benjamin, S., Soccol, C., Nigam, P., Krieger, N., and Soccol, V., 1999. The realm of microbial lipases in biotechnology: a review. Biotechnology and Applied Biochemistry, 29: 119–131. Patel, M. T., Nagarajan, R., and Kilara, A., 1996. Hydrolysis of milk fat by lipase in solventfree phospholipids reverse micellar media. Journal of Food Science, 61 (1): 33-39. Pleiss, J., 2000. Molecular basis of specificity and stereoselectivity of microbial lipases toward triacylglycerols. In: Enzymes in Lipid Modification. (eds.) Bornscheuer, U., Wiley-VCH. Weinheim, pp: 85-99. Qian, J. H., and Xu, J. H., 2004. Catalytic performance of a highly enantioselective (R)-ester hydrolase from a new isolate Acinetobacter sp. CGMCC 0789. Journal of Molecular Catalysis B: Enzymatic, 27: 227–32. Rahman, R. N. Z., Chin, J. H., Salleh, A. B., and Basri, M., 2003. Cloning and expression of a novel lipase gene from Bacillus sphaericus 205y. Molecular Genetics and Genomics, 269: 252-260. Rahman, R. N. Z., Leow, T. C., Basri, M., and Salleh, A. B., 2005. Secretory expression of thermostable T1 lipase through Bacteriocin Release Protein (BRP). Protein Expression and Purification, 40 (2): 411-416. Ruiz, C., Blance, A., Pastor, F. I. J., and Diaz, P., 2002. Analysis of Bacillus megaterium lipolytic system and cloning of LipA, a novel subfamily I.4 bacterial lipase. FEMS Microbiology Letters, 10746: 1-5.

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Sakiyama, T., Yoshimi, T., Miyake, A., Umeoka, M., Tanaka, A., Ozaki, S., and Nakanishi, K., 2001. Purification and characterization of a monoacylglycerol lipase from Pseudomonas sp. LP7315. Journal of Bioscience and Bioengineering, 91 (1): 27-32. Saxena, R. K., Ghosh, P. K., Gupta, R., Sheda Dvidson, W., Bradoo, S., and Gulati, R., 1999. Microbial lipases, potential biocatalysts for the future industry. Current Science, 77: 101115. Schmidt-Dannert, C., 1999. Recombinant microbial lipases for biotechnological applications. Bioorganic and Medicinal Chemistry, 7: 2123-2130. Schmidt-Dannert, C., Rúa, M. L., Atomi, H., and Schmid, R. D., 1996. Thermoalkalophilic lipase of Bacillus thermocatenulatus. I. Molecular cloning, nucleotide sequence, purification and some properties. Biochimica et Biophysica Acta, 1301: 105-114. Schmidt-Dannert, C., Sztajer, H., Stöcklein, W., Menge, U., and Schmid, R. D., 1994. Screening, purification and properties of a thermophilic lipase from Bacillus thermocatenulatus. Biochimica et Biophysica Acta, 1214: 45-53. Sharma, R., Soni, S. K., Vohra, R. M., Gupta, L. K., and Gupta, J. K., 2002. Purification and characterization of a thermostable alkaline lipase from a new thermophilic Bacillus sp. RSJ-1. Process Biochemistry, 37: 1075-1084. Shen, D., Xu, J. H., Gong, P. F., Wu, G. H., and Liu, Y. Y., 2000. Isolation of an esterase producing Trichosporon brassicae and its catalytic performance in kinetic resolution of ketoprofen. Canadian Journal of Microbiology, 47: 1101–6. Shiomori, K., Ishimura, M., Baba, Y., Kawano, Y., Kuboi, R., and Komasawa, I., 1996. Characteristics and kinetics of lipase-catalysed hydrolysis of olive oil in a reverse micellar system. Journal of Fermentation and Bioengineering, 81 (2): 143-147. Sigurgísladóttir, S., Konráðsdóttir, M., Jónsson, A., Kristjánsson, J. K., and Matthiasson, E., 1993. Lipase activity of thermophilic bacteria from icelandic hot springs. Biotechnology Letters, 15 (4): 361-366. Sinchaikul, S., Sookkheo, B., Phutrakul, S., and Pan, F. M., 2001. Optimization of a thermostable lipase from Bacillus stearothermophilus P1: Overexpression, purification, and characterization. Protein Expression and Purification, 22: 388-398. Snellman, E. A., Sullivan, E. R., and Colwell, R. R., 2002. Purification and properties of the extracellular lipase, LipA, of Acinetobacter sp. RAG-1. The FEBS Journal, 269: 5771– 5779. Sonnet, P. E., and Gazzillo, J. A., 1991. Evaluation of lipase selectivity of hydrolysis. Journal of the American Oil Chemist’s Society, 68: 11-15. Soumanou, M. M., Bornscheuer, U. T., Schmid, U., and Schmid, R.D., 1999. Crucial role of support and water activity on the lipase-catalyzed synthesis of structured triglycerides. Biocatalysis Biotransformations, 16: 443-459. Sugihara, A., Tani, T., and Tominaga, Y., 1991. Purification and characterization of a novel thermostable lipase from Bacillus sp. Journal of Biochemistry, 109 (2): 211-215. Sugihara, A., Ueshima, M., Shimada, Y., Tsunasawa, S., and Tominaga, Y., 1992. Purification and characterization of a novel thermostable lipase from Pseudomonas cepacia. Journal of Biochemistry, 112: 598-603. Sugihara, A., Shimada Y., and Tominaga, Y., 1988. Purification and characterization of Asperigillus niger lipase. Agricultural and Biological Chemistry, 52(6): 1591-1592.

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Sugiura, M., and Isobe, M., 1974. Studies on the lipase of Chromobacterium viscosum. 3. Purification of a low molecular weight lipase and its enzymatic properties. Biochimica et Biophysica Acta, 341: 195-200. Takamoto, T., Shirasaka, H., Uyama, H., and Kobayashi, S., 2001. Lipase-catalyzed hydrolytic degradation of polyurethane in organic solvent. Chemistry Letters, 6: 492–3. Tan, Y., and Miller, K. J., 1992. Cloning, expression and nucleotide sequence of a lipase gene from Pseudomonas fluorescens B52. Applied and Environmental Microbiology, 58 (4): 1402-1407. Tanaka, K., Yoshida, K., Sasaki, C., and Osano, Y. T., 2002. Practical asymmetric synthesis of the herbicide (S)-indanofan via lipase-catalysed kinetic resolution of a diol and stereoselective acid-catalysed hydrolysis of a chiral epoxide. The Journal of Organic Chemistry, 67: 3131-3133. Thomson, C. A., Delaquis, P. J., and Mazza, G., 1999. Detection and measurement of microbial lipase activity: A review. Critical Reviews in Food Science and Nutrition, 39 (2): 165-187. Undurraga, D., Markovits, A., and Erazo, S., 2001. Cocoa butter equivalent through enzymic interesterification of palm oil midfaction. Process Biochemistry, 36: 933-939. Verger, R., 1997. Interfacial activation of lipases: facts and artifacts. Trends in Biotechnology, 15: 32-38. Villeneuve, P., Muderhwa, J. M., Graille, J., and Haas, M. J., 2000. Customizing lipases for biocatalysis: a survey of chemical, physical and molecular biological approaches. Journal of Molecular Catalysis B: Enzymatic, 9: 113-148. Vujaklija, D., Schröder, W., Abramić, M., Zou, P., Leščić, I., Franke, P., and Pigac, J., 2002. A novel streptomyces lipase: cloning, sequencing and high-level expression of the Streptomyces rimosus GDS(L)-lipase gene. Archives of Microbiology, 178: 124-130. Vulfson, E. N., 1994. Lipases: their Structure, Biochemistry and Application. (eds.) Woolley, P., and Petersen, S. B., Cambridge University Press. United Kingdom, pp: 271–288. Wang, Y., Srivastava, K. C., Shen, G. J., and Wang, H. Y., 1995. Thermostable alkaline lipase from a newly isolated thermophilic Bacillus, strain A30-1 (ATCC 53841). Journal of Fermentation and Bioengineering, 79 (5): 433-438. Wang,Y, Srivastava, K. C., Shen, G. J., and Wang, H. Y., 1995. Thermostable alkaline lipase from a newly isolated thermophilic Bacillus strain, A30-1 (ATCC 53841). Journal of Fermentation and Bioengineering, 79: 433–438. Weber, N., Klein, E., and Mukerjee, K. D., 1999. Long chain acyl thioesters prepared by solvent free thioesterification and transesterification catalyzed by microbial lipases. Applied Microbiology and Biotechnology, 51: 401–4. Wohlfarth, S., Hoesche, C., Strunk, C., and Winkler, U., 1991. Molecular genetics of the extracellular lipase of Pseudomonas aeruginosa PAO1. Journal of General Microbiology, 138: 1325-1335. Yan, F., Yong-Goe, J., Kazuhiko, I., Hiroyasu, I., Susumu, A., Tohru, Y., Hiroshi, N., Shugui, C., Ikuo, M., and Yoshitsugu, K., 2000. Thermophilic phospholipase A2 in the cytosolic fraction from the archaeon Pyrococcus horikoshii. Journal of the American Oil Chemists' Society, 77: 1075–1084. Yeoh, H. H., Wong, F. M., and Lin, G., 1986. Screening for fungal lipases using chromogenic lipid substrates. Mycologia, 78: 298–300.

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Zhang, L. Q., Zhang, Y. D., Xu, L., Li, X. L., Yang, X. C., Xu, G. L., Wu, X. X., Gao, H. Y., Du, W. B., Zhang, X. T., and Zhang, X. Z., 2001. Lipase catalyzed synthesis of RGD diamide in aqueous water-miscible organic solvents. Enzyme and Microbial Technology, 29: 129–35.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 23-39

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 2

PROTEASE: INTRODUCTION Abu Bakar Salleh, Che Nyonya Abdul Razak, Raja Noor Zaliha Raja Abd. Rahman and Mahiran Basri ABSTRACT Protease can be obtained from plants, animals and microorganisms. They are grouped based on their specificities or mechanism of action. Proteases have wide industrial applications, mainly in food industry, detergent, medical and biotechnology. Many microbial proteases have been purified, characterized and commercialized. Microbial proteases are the most frequently utilized commercial enzymes with wide application in various industries such as detergent, textile, leather and food industry. Proteases comprise 60% of total enzyme sales, with detergent enzyme being the major protease user.

INTRODUCTION Protease is a group of enzymes that catalyse hydrolysis of peptide bonds in protein. The main function of the extracellular proteases are similar to many other polymers degrading extracellular enzymes in nutrition. The enzymes hydrolyze large polypeptides or proteins to smaller materials that can be absorbed by the cells. Apart from that, owing to their hydrolytic ability, intracellular proteases play an important role in many biological processes such as in the utilization of proteinaceous nutrient, release of protein, zymogen activation, autolysis, spore germination and other physiological phenomena. The proteolytic reaction of proteases degrade the protein into peptone, polypeptides, dipeptides and finally into amino acids (Cappuccino and Sherman, 1992). Proteases are also involved in protein degradation to amino acids and peptides as nutrition source and in protein mobilization, post translation reactions, modulation of gene expression, enzyme modification and secretion of many protein enzymes (Suhartono et al., 1997).

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Proteases have wide industrial applications, mainly in food industry, detergent, medical and biotechnology. Now many microbial proteases have been purified, characterized and commercialized. In food industry, application of protease includes in production of cheese, solubilisation of fish paste and softening of meat. Apart from that, proteases are also widely used in the leather industry. From the economic and technological aspects, microbial proteases are the most frequently utilized commercial enzyme with wide application in various industries such as detergent, leather, silk, bread, meat and brewery. Proteases comprise 60% of total enzyme sales, with detergent enzyme being the major protease user. Proteases from Bacillus sp. contribute to 25% sale (Jasvir et al., 1999). For example, Bacillus licheniformis strain is listed in the third edition of Food Chemicals Codex as a source of carbohydrate enzyme and protease used in food processing. Important detergent proteases in the market includes subtilisin Carlsberg, subtilisin BPN’, Alcalase, Esparase and Savinase (Gupta et al., 1999). Alkaline proteases in wash detergent help to release protein dirt such as blood and food. Wide diversity of proteases contrast with their narrow specificity has attracted world attention in exploiting then for physiological and biotechnological uses (Fox et al., 1991; Poldermans 1990). Major world protease producers are listed in Table 1. Table 1. Major protease producers Companies’ Country

Market share (%)

Novo Industries Denmark Gist-Brocades Netherlands Genencor International United States Miles Laboratories United States Others

40 20 10 10 20

(Source: Rao et al., 1998)

SOURCES Protease can be obtained from plants, animals and microorganisms. Utilization of plants as a source for proteases depends on a number of factors such as availability of land for farming, suitable temperature and it is time consuming. Proteases produced from plants include papain, bromelin, keratinize and ficin. Papain is produced from Carica papaya. Bromelain is from the stalk and fruit of pineapples and ficin from the latex of Ficus glabrata fig tree. All three proteases have similar specificity although obtained from different sources. Some botanical plant groups produce keratinase which can degrade hair. Digestion of hair and fur is important in production of essential amino acids and to avoid blockage of effluent systems (Rao et al., 1998). Examples of proteases produced from animals are protaminase, trypsin, chymotrypsin, pepsin and renin. Protaminase is isolated from bovine pancreas, sea lions and pigs, while

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trypsin and chymotrypsin, both known as proteinase from pancreas, are produced by acinar cells of pancreas and secreted in the form of zymogen each known as trypsinogen and chymotrypsinogen. Pure chymotrypsin is expensive and used in diagnostic and analytical applications. It is specific in hydrolyzing peptide bond where carboxyl group is provided by one of the three aromatic amino acids i.e. phenylalanine, tyrosine or tryptophan. It can be used in deallergenizing milk protein hydrolysis (Rao et al., 1998). Pepsin (E.C. 3.4.4.1) can be obtained from gastric mucosal cells of pig and renin (E.C. 3.4.4.3) from lamb stomach. Active enzyme is released from its zymogen, and pepsinogen through autocatalysis in the presence of hydrochloric acid. It catalyses the hydrolysis of peptide bond between two hydrophobic amino acids (Rao et al., 1998). Studies on intracellular proteases are rather scarce. Among microbes reported to be intracellular protease producers include Clostridium perferingens tip a (Park and Labbe, 1990) and Streptococcus saliarius subsp. thermophilus (Tsakalidou and Kalantzopolus, 1992). Protease that binds to membrane is the least studied protease. It was reported to be produced by Escherichia coli (Palmer and St. John, 1987) and Lactococcus lactis subsp. cremoris AMI (Visser et al., 1991).

TYPES AND PROPERTIES The International Union of Biochemistry and Molecular Biology (1984) suggested the use of the term peptidase for subset of peptide bond hydrolases (Subclass E.C 3.4.). The commonly used term of protease is synonymous with peptidase. Peptidases comprise two groups: endopeptidases and exopeptidases, which cleave peptide bond of protein and eliminating amino acid sequentially from N or C terminal. The term proteinase can also be used for endopeptidase. The four mechanistic classes of proteinases recognized by IUBMB are as listed below. Modern nomenclature scheme is:

Peptidases (or proteases) (E. C 3.4.-.-)

Exopeptidases (E. C 3.4.11-19.-)

Endopeptidases (or proteinases) (E. C 3.4.21-99.-)

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A. B. Salleh, C. N. A. Razak, R. N. Z. R. A. Rahman et al. Exopeptidases can be categorized into a number group: a. Aminopeptidases (3.4. 11-19) which hydrolyze one residue from -N terminal b. Carboxypeptidases (3.4.16-17) which hydrolyse one residue from -C terminal c. Dipeptidases (3.4.13) which are specific for dipeptide, including enzymes that cleave dipeptide unit from either N terminal (3.4.14) or C terminal (3.4.15). d. Omega peptidases (3.4.19) are the enzymes which hydrolyse amino acid residue substituted from either N terminal (3.4.14) or C terminal (3.4.15).

Proteinase groups include enzyme endopeptidases and peptidyl-peptide hydrolases. Hartley (1960) has divided the proteinases into four groups based on reaction mechanism, namely serine proteinase, (3.4.21), thiol proteinase (3.4.22), acid proteinase (3.4.23) and metallo proteinase. Even now with development in chemical study particularly in the enzyme active site, this classification concept is still valid and lasting, however the name of 2 groups have been changed from, thiol to cysteine proteinase and acid to aspartic proteinase. Classification based on catalytic type was suggested to classify the group which had evolutionary linkages to proteases (Rawlings and Barrett, 1993). This classification can be accessed from SwissProt database. Apart from the four mechanistic classes, there is one section in enzyme nomenclature for protease with unknown reaction mechanism. It indicates that this reaction mechanism has not been elucidated as such there may be other novel proteases. Reaction mechanism of protease has become a field of study that attracts many researchers. Catalytic site of protease is sandwiched on one or both sides by specificity subsite, each one capable to accept the side chain of amino acid residue of the substrate. This site is numbered from site catalytic S1 via Sn towards the N terminus of the structure and S1' via Sn' towards C terminus. Residue accepted from the substrate is assigned as Pl via Pn and P1' via Pn' (Figure 1). Protease

: N Sn---- S3—S2—S1* S’1—S’2—S’3---- S’n C

Substrate

: N Pn-----P3—P2---P1--- P’1—P’2—P’3----P’n C

Figure 1: Active sites of protease. catalytic site of protease marked by * and scissile bond shown by; S1 via Sn and S1' via Sn' are specificity subsites of the enzyme, while P1 via Pn and P1' via Pn' are substrate residues accepted by enzyme subsites (Rao et al., 1998).

Serine Protease (EC 3.4.21) Serine protease comprises two subfamilies, superfamily chymotrypsin and subtilisin (Hourses 1976). Chymotrypsin group including mammalian digestive protease, has structural homology with trypsin, elastase and thrombin (Graycar, 1999). Proteases related to chymotrypsin can be obtained from microorganisms, plants and vertebrate animals, whereas proteases related to subtilisin can only be obtained from bacteria. These proteases hydrolyse peptide bond where the carboxyl group is contributed by the lysine and arginine residue (Table 2). General 3D structure is different in both groups but they

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have similar active site geometry and reaction occurs by similar mechanism. Serine proteinases exhibit different specificity related to different amino acid substitutions in the enzyme subsites (see nomenclature of Schechter and Berger, 1967) reacting with substrate residue. Some enzymes have broad interactive site with the substrate while others have limited specificity to P1 substrate residue. Three residues that contribute to the catalytic triad are essential in the catalytic process (i.e. His 57, Asp 102 and Ser 195 chymotrypsin numbering). The first step in the catalysis is the production of acyl enzyme intermediate between substrate and Serine. It is necessary to produce this covalent intermediate via a negatively charged tetrahedral intermediate, and subsequently the cleavage of the peptide bond. During the second step or deacylation, the acyl-enzyme is hydrolysed by a water molecule to release the peptide and to restore the Ser-hydroxyl of the enzyme. The deacylation also involves the formation of a tetrahedral transition state, where it proceeds via the reverse reaction pathway of acylation. A water molecule is attacking the nucleophile instead of the Ser residue. The His residue provides a base that accepts the OH group of the reactive Ser. Table 2. Specificity of proteases* Enzyme Trypsin Chymotrypsin, subtilisin Staphylococcus V8 protease Papain Thermolysin Pepsin a *

Peptide bond cleaveda -Lys (or Arg) ↓-----Trp (or Tyr, Phe, Leu)↓ ------Asp (or Glu) ↓ ------Phe (or Val, Leu)-Xaa↓--------↓Leu (or Phe) ------Phe (or Tyr, Leu) ↓ - Trp (or Phe, Tyr)

The arrow indicates the site of action of the protease. Xaa, any amino acid residue. (Source: Rao et al., 1998)

Serine protease is widely found in many varieties, where more than 40% of microbial proteases reported are from this group of enzyme. It is also known as alkaline protease referring to their optimum pH in the range of pH 9 to 13. Alkaline serine protease is widely used in detergent. It is the most extensively used microbial enzyme in the industrial sector. In fact, subtilisin Novo (BPN’) and subtilisin Carlsberg, two main enzymes in this group were the first enzymes produced industrially. A lot of information is available on the two enzymes as they have industrial potential, accordingly they have become the main target for protein engineering. Amino acid sequence for both enzymes has long been known (Markldan and Smith, 1967; Smith et al., 1968). Three-dimensional structure for subtilisin BPN’ has also been obtained since 1969 while the structure of subtilisin Carlsberg was reported later (Bode et al., 1987; Neidhart and Petsko, 1988). Serine protease can be classified into three main types based on substrate specificity; trypsin-like enzyme, chymotrypsin-like enzyme and elastase-like enzyme (Polgar, 1987). Trypsin-like enzyme will cleave substrates containing positively-charged amino acid such as lysine and arginine, whereas elastase-like enzyme will select substrates with small aliphatic side chains and chymotrypsin-like enzyme prefers substrates containing large aromatic and aliphatic side chains.

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Serine protease usually has a molecular weight of 18 and 35 kDa (Rao et al., 1998). It requires an activator, although Ca2+ ion is required for partial activation of the proenzyme and also for enzyme’s stability. In 1964, Gold and Fahrney reported phenylmethane sulfonyl fluoride (PMSF) could deactivate trypsin and chymotrypsin and since then, sulfonyl flouride compounds are widely used as inhibitors for serine protease. Now there are other inhibitors that inhibit serine protease specifically, for example 4,amidinophenylmethane sulfonyl fluoride (APMSF), leupeptin and antipain which is a specific inhibitor for trypsin-like enzymes (Laura et. al., 1980; Umezawa and Aoyagi, 1977) Meanwhile, the most important inhibitors of chymotrypsin-like protease are N-tosyl-L-lysine chloromethyl ketone and soyabean trypsin inhibitor (Gupta et al., 2002). Whereas elastinal is a specific inhibitor for elastase (Robert et al., 1997).

Cysteine Protease (EC 3.4.22) This family of enzymes includes the plant protease such as papain, actinidin or bromelain, several mammalian lysosomal cathepsins, the cytosolic calpains (calciumactivated) as well as several parasitic proteases (e.g Trypanosoma, Schistosoma). Papain is the archetype and the best studied member of the family. Recent elucidation of the x-ray structure of Interleukin-1-beta Converting Enzyme has revealed a novel type of fold for cysteine proteinases. Similar to the serine proteinases, catalysis proceeds via the formation of a covalent intermediate involving a cysteine and a histidine residue. The essential Cys25 and His159 (papain numbering) play the same role as Ser195 and His57 respectively. The nucleophile is a thiolate ion rather than a hydroxyl group. The thiolate ion is stabilized via the formation of an ion pair with the neighbouring imidazolium group of His159. The attacking nucleophile is the thiolate-imidazolium ion pair in both steps and a water molecule is not required. Cysteine protease was known as thiol protease as its catalyst activity depends on the presence of a thiol group in the enzyme molecule. This cysteine protease contains one cysteine sulphydryl group (Cys25) and histidine imidazole group (His159) at the enzyme active site (Polgar and Halasz, 1982). Cysteine protease is isolated from various sources including animals (cathepsin B) (Guicciardi et al., 2001), plants (bromelain, ficin, papain) (Oliva et al., 2004) and microbes (clostripain, Plasmodium falciparum cysteine protease (Rosenthal, 2004). Most cysteine protease has molecular weights between 20,000 to 50,000 Da (Rao et al., 1998) and is active at neutral pH. Various compounds were reported to be inhibitors for this enzyme, including iodo- and bromo acetate, p-chloromercuric benzoate, Nethylmaleimide, where all these compounds cause the alkylation of the cysteine sulphydryl group resulting in the activation of enzyme.

Aspartic Protease (EC. 3. 4. 23) Most of the aspartic proteinase enzymes belong to the pepsin family. The pepsin family includes digestive enzymes such as pepsin and chymosin as well as lysosomal cathepsins D and processing enzyme such as renin, and certain fungal proteases (penicillopepsin, rhizopuspepsin, and endothiapepsin). A second family comprises viral proteinases such as the

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29

protease from the HIV virus also called retropepsin. Crystallographic studies have shown that these enzymes are bilobed molecules with the active site located between two homologous lobes. Each lobe contributes one aspartate residue of the catalytically active diad of aspartates. These two aspartyl residues are in close geometric proximity in the active molecule and one aspartate is ionized, whereas the second one is unionized at the optimum pH range of 2-3. Retropepsins, are monomeric, with only one catalytic aspartate and dimerization is required to form an active enzyme (Rao et al., 1998). In contrast to serine and cysteine proteases, catalysis by aspartic proteinases does not involve a covalent bond in between though a tetrahedral intermediate exists. The nucleophilic attack is achieved by two simultaneous proton transfer; one from a water molecule to the diad of the two carboxyl group and a second one from the diad to the carbonyl oxygen of the substrate with the concurrent CO-NH bond cleavage. This general acid-base catalysis, which may be called a "push-pull" mechanism leads to the formation of a non covalent neutral tetrahedral intermediate (Rao et al., 1998). Aspartic protease is limited to the eukaryotic cells only. It was known as acidic protease as this protease has an optimum pH in the range of pH 3.5 to 5.5. Generally, aspartic protease acts on peptide bond between large hydrophobic amino acid residues. Many microbial aspartic proteases have been crystallized from Mucor pusillus (Newman et al., 1993), Candida albicans (Abad-Zapatero et al., 1996) and Rhizomucor miehei (Yang et al., 1997). Molecular weight for aspartic protease is in the range of 30,000 to 45,000 Da (Rao et al.,1998). The first inhibitor identified for this protease is pepstatin A (Umezawa, 1982; Fitzgerald et al., 1990). They are also sensitive to diazoketone compounds such as diazoacetyl-DL-norleucine methyl ester (DAN) and 1,2-epoxy-3-(p-nitrophenoxy)propane (EPNP) in the presence of copper ions (Rao et al., 1998).

Metallo Protease (EC 3.4.24) The metallo proteinases may be one of the older classes of proteinases found in bacteria, fungi as well as in higher organisms. They differ widely in their sequences and their structures but the great majority of enzyme contains a zinc atom which is catalytically active. In some cases, zinc may be replaced by another metal such as cobalt or nickel without loss of activity. Bacterial thermolysin has been well characterized and its crystallographic structure indicates that zinc is bound by two histidines and one glutamic acid. Many enzymes contain the sequence HEXXH, which provides two histidine ligands for the zinc whereas the third ligand either a glutamic acid (thermolysin, neprilysin, alanyl aminopeptidase) or a histidine (astacin). Other families exhibit a distinct mode of binding of the Zn atom. The catalytic mechanism leads to the formation of a non covalent tetrahedral intermediate after the attack of a zinc-bound water molecule on the carbonyl group of the scissile bond. This intermediate is further decomposed by transfer of the glutamic acid proton to the leaving group. This type of protease is widely found in bacteria, fungi and higher organism. Thermolysin from Bacillus thermoproteoliyticus is one of the most studied enzymes from this group. The characteristics of the enzyme are known in detail, with the amino acid sequence and its crystallographic structure has been elucidated (Colman et al., 1972). Metallo protease is specific to peptide bond between two amino acid residues with non polar side chain and bulky amino acid such as phenylalanine and leucine (Barrett, 1986). Normally the optimum

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activity for metallo protease is in the pH range between pH 7.0 to 8.0. This protease group is the most unstable group compared to other types of protease and will undergo autolysis process quickly in the pH range of above pH 9.0 and below pH 6.0. Inhibitors usually identified as metallo protease inhibitors are ethylenediaminetetraacetic acid (EDTA), 1,10phenanthroline and 2,2-bipyridine (2,2’ dipyridyl). But EDTA is a good chelating agent for calcium, and is not a specific inhibitor for metallo protease as it can inhibit other enzymes that require calcium for their activity such as serine protease. However, 1,10-phenanthroline is more specific as an inhibitor for metallo protease (Powers and Harper, 1986). Some peptide inhibitors from microbes specific for metallo protease were reported including phoshoramidon (N[O←L-rhamnopyranosylhydroxyphosphinyl]-L-leucyl-Ltryptophan), bestatin and amastatin (Suda et al., 1973; Umezawa, 1982). Phoshoramidon peptide inhibitor produced by actinomycetes is the first inhibitor shown to be specific and potent for neutral metallo protease such as thermolysin (Suda et al., 1973). In phosphoramidon molecule, there is a tetra N-phosphoryl moiety that can interact with the metal in the enzyme active site of metallo protease.

INDUSTRIAL APPLICATIONS OF PROTEASES Detergent Proteases are one of the standard ingredients of all types of detergents ranging from those used for household laundering to reagents used for cleaning contact lenses or dentures. The use of protease in laundry detergents accounts for approximately 25% of the total worldwide sales of enzyme. The use of microbial proteases as an active agent in detergent started in 1960’s. The first detergent containing the bacterial enzyme was introduced in 1956 under the trade name BIO-40. In 1960, Novo Industry A/S introduced alcalase, produced by Bacillus licheniformis; with commercial name of BIOTEX. This was followed by Maxatase, a detergent made by Gist-Brocades (Rao et al., 1998). Now microbial proteases are found in most types of detergent brands marketed. The increased usage of these proteases as detergent additives is mainly due to the cleaning capabilities of these enzymes in environmentally acceptable, nonphosphate detergents. In addition to improve washing efficiency, the use of enzymes allows lower wash temperatures and shorter periods of agitation, often after a preliminary period of soaking (Kumar and Takagi, 1999). Ideally, proteases and other enzymes used in detergent formulations should have high activity and stability over a broad range of pH and temperature. The enzymes used should be effective at low levels (0.4–0.8%) and should also be compatible with various detergent components along with oxidizing and sequestering agents. They must also have a long shelf life. Recently, proteases with such properties from Vibrio fluvialis (Venugopal and Saramma, 2006) and Nocardiopsis sp. (Moreira et al., 2002) have been reported. The key parameter for the best performance of a protease in a detergent is its pI. It is known that a protease is most suitable for this application if its pI coincides with the pH of the detergent solution. Esperase and Savinase T (Novo Industry), produced by alkalophilic Bacillus spp., are two commercial preparations with very high isoelectric points (pI 11); hence, they can withstand higher pH ranges. Due to the present energy crisis and the awareness for energy conservation, it is desirable to use protease that is

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active at lower temperatures. A combination of lipase, amylase, and cellulase is expected to enhance the performance of protease in laundry detergents (Rao et al., 1998). The biggest market for detergents is in the laundry industry, amounting to a worldwide production of 13 billion tons per year. Among the factors that contribute towards the use of protease enzyme in detergent is the consumer environmental awareness that prefers washing process to be done at lower temperature and to avoid use of compound containing phosphate that can contaminate the environment (Rao et al., 1998). Early detergent formulation was developed for washing at high temperature and usually contained sodium phosphate, tribasic and sodium perborate. Sodium phosphate was used as the effective dirt removal agent, while sodium perborate as a whitening agent active at temperature above 60oC. However man has become more sensitive to environmental pollution by phosphate and the increase use of polyester, so the use of these two compounds were reduced or completely eliminated. Consequently, detergent quality has become inferior compared to the old detergent. However, when enzymes, including protease, are used as one of the detergent components, it enhanced the effectiveness of the new detergent formulation. Current detergent formulations not only able to wash at low or moderate temperature but the components are also environmental friendly (Rao et al., 1998). Another factor that contributes towards the increase in the use of enzyme in detergent is the higher production of potential enzymes. Now R&D groups are working hard, competing to acquire better proteases that are able to remove stains and are more stable. Enzyme suppliers continuously screen microbes that produce proteases with greater potential and are more stable. At the same time, new surfactant compounds, ‘builders’ and ‘extenders’ are being used, and the industry is forced to screen for proteases that are stable in the new detergent component. Apart from that, with the introduction of detergent in liquid form, enzyme suppliers also face another problem in trying to enhance the stability of enzyme in aqueous environment, as normally the half lives of protease are longer for enzyme in the powder or granular form as compared to the liquid form (Rao et al., 1998). Conventionally, detergents have been used at elevated washing temperatures, but at present there is considerable interest in the identification of alkaline proteases which are effective over a wide temperature range (Oberoi et al., 2001). One of the enzymes that meets the requirement is alkaline serine protease, which is not inhibited at low temperature but like other enzymes, unfortunately, their rate of catalysis is reduced at low temperature.Therefore, this has pushed enzyme manufacturers to look for novel enzyme that can act under low temperatures. In Japan, Novo Nordisk Bioindustry has developed a detergent protease called Kannase, which keeps its high efficiency, at even very low temperatures (10-20°C) (Gupta et al., 2002). As the environment for microbial protease in detergent is harsh, most enzymes used are from alkaline serine protease isolated from alkalophilic Bacillus sp. Protease produced by this group of bacteria can survive in alkaline environment and high temperature. Subtilisin Carlsberg produced by B. licheniformis was the first microbial protease used in detergent (Gupta et al., 1999). In a recent study by Gupta et al. (2002), the wash performance analysis of a SDS-stable alkaline protease from Bacillus sp. RGR-14 (Oberoi et al., 2001) has shown an increase in reflectance (14% with grass stains, 25% with blood stain) after enzyme treatment. Currently, many microbial proteases have been patented to be used in detergent formulation but they are still not available in the market. All detergent proteases currently used in the market are serine proteases produced by Bacillus strains. Fungal alkaline protease

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is advantageous due to the ease of downstream processing to prepare a microbe-free enzyme. An alkaline protease from Conidiobolus coronatus was found to be compatible in commercial detergents used in India (Phadatare et al., 1993) and retained 43% of its activity at 50°C for 50 min in the presence of Ca2+ (25 mM) and glycine (1 M) (Bhosale et al., 1995).

Cheese Production The major application of protease in the dairy industry is in the manufacture of cheese. The milk-coagulating enzyme falls into three main categories, animal rennets, microbial milk coagulants, and genetically engineered chymosin. Both animal and microbial milkcoagulating proteases belong to a class of aspartate protease that has molecular weights from 30,000 to 40,000 Da. Rennet extracted from the fourth stomach of unweaned calves contains the highest ratio of chymosin (EC 3.4.23.4) to pepsin activity. Renin, a protease enzyme (chymosin) from calves has long been used for formation of clot or to coagulate milk in cheese production industry (Rao et al., 1998). However, the cost is high as the source of the enzyme is difficult to obtain. The cheese industry has attempted using pepsin from bovine or pigs as alternatives as pepsin has very high proteolytic activity. However, followed by the ban by some religious groups on the use of rennet from animal sources, microbial rennet was accepted as the alternative enzyme in the 1970’s (Rao et al., 1998). A world shortage of calf rennet due to the increased demand for cheese production has intensified the search for alternative microbial milk coagulants. The microbial enzyme exhibited two major drawbacks, (i) the presence of high levels of nonspecific and heat-stable protease, which led to the development of bitterness in cheese after storage; and (ii) a poor yield. Due to this, a few acid proteases have been isolated from microbes that were used in cheese production, including acid protease from Mucor meihei and Irpex lactis (Godfrey and West, 1996). Even though the use of microbial rennet has increased from 10% in 1978 to 19% in 1986 (Aunstrup, 1980; Hepner and Male, 1986), renin from bovine calves is still preferred by cheese manufacturers. Microbial rennet has been chemically modified to enhance its thermal stability, although this modification has positive effect, it also increases the proteolysis rate during pasteurisation and in the process causes the milk to boil (Rao et al., 1998). The current role of industry is to obtain microbial rennet that is closest to the ability of calf renin. Although effort has been put by the industry to clone calf chymosin gene into microbes with some degree of success, the use of clones is still not adopted. The delay is probably due to the concern of the consumers for product of cloned microbes. It is hoped that the product would be in the market for the future. Extensive research in this area has resulted in the production of enzyme that is completely inactivated at normal pasteurization temperatures and contains very low levels of nonspecific protease. In cheese making, the primary function of protease is to hydrolyze the specific peptide bond (Phe105-Met106 bond) to generate para-casein and macropeptides. Chymosin is preferred due to its high specificity for casein, which is responsible for its excellent performance in cheese making. The protease produced by GRAS (genetically regarded as safe)-cleared microbes such as M. meihei, Bacillus subtilis, and Endothia parasitica are gradually replacing chymosin in cheese making. In 1988, chymosin produced through recombinant DNA technology was first introduced to cheesemakers for evaluation. Genencor

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International increased the production of chymosin in Aspergillus niger var. awamori to commercial levels. Currently, there are three recombinant chymosin products available and awaiting legislative approvals for their use in cheese making (Godfrey and West, 1996).

Leather Industry Leather processing involves several steps such as soaking, dehairing, ‘bating’, and tanning. The major building blocks of skin and hair are proteinaceous. The conventional methods of leather processing involve hazardous chemicals such as sodium sulphide, which create problems of pollution and effluent disposal. The use of enzyme as alternatives to chemicals has proven successful in improving leather quality and in reducing environmental pollution. In leather industry, there are 2 processes involving the use of enzyme protease; hair removal process from leather and ‘bating’. Proteases are used for selective hydrolysis of noncollagenous constituents of the skin and for removal of nonfibrillar proteins such as albumins and globulins. The purpose of soaking is to swell the hide. In hair removal process, the usual technique is to use chalk and sodium sulphide whereby the leather would swell, becoming loose and destroying the hair (Rao et al., 1998).This method is cheap and fast. However, the pressure against using environmentally unfriendly compounds and also the health of workers that are forced to inhale hydrogen sulphide gas has forced the leather industry to use protease from microbial or transfer the industry to another country that has relaxed regulation of pollution. Alkaline proteases with elastolytic and keratinolytic activitites can be used in leather-processing industries (Gupta et al., 2002). Alkaline proteases will speed up the process of dehairing, because the the alkaline conditions anable the swelling of hair roots; and the subsequent attack of protease on hair follicle protein allows easy removal of the hair. Therefore, most entrepreneurs adopt the second alternative as it is cheaper. However, the first alternative introduces the use of microbial protease in leather industry. Alkaline serine proteases isolated from alkalophilic Bacilli is mostly used in this process. In this enzymatic process, leather is treated with 8% of chalk and 0.1% of enzyme for 18-24 hours at temperature of 20-30ºC, pH 12-12.5. This process will destroy unwanted leather pigment, leather surfaced is increased, fine hair is removed and ‘bating’ process can be shortened or not required at all (Rao et al., 1998). ‘Bating’ process of leather is a process which makes leather more malleable and elastic. The bating following the dehairing process involves the degradation of elastin and keratin, removal of hair residues, and the deswelling of collagen, which produces a good, soft leather mainly used for making leather clothes and goods. In addition, studies carried out by different workers have demonstrated the successful use of alkaline proteases in leather tanning from Aspergillus flavus (Malathi and Chakraborty, 1991), Streptomyces sp. (Mukhopadhyay and Chandra, 1993) Bacillus amyloliquefaciens (George et al., 1995) and B. subtilis (Varela et al., 1997) Leather used in the glove production, for example, has extensive hair ‘bating’ process to enable it to be formed. Different from hair removal process, ‘bating’ process is traditionally an enzymatic process. But the enzyme used comprising trypsin or crude enzyme such as pancreatin. Serine protease from bacteria was used to replace trypsin from animals but without success because of its high proteolytic activity (Rao et al., 1998). Enzyme can be

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directly used on the leather or mixed with wood powder and if the leather is dry, enzyme can be mixed into the washing solution. Novo Nordisk manufactures three different proteases, Aquaderm, NUE, and Pyrase for use in soaking, dehairing, and bating, respectively (Rao et al., 1998). In addition, Sana et al. (2006) is also investigating a single salt, solvent, detergent and bleach tolerant alkaline serine protease produced from a new gamma-Proteobacterium for a commercial application during the dehairing process in the leather industry.

Bread Production Enzymes such as amylases, proteases and xylanases can be used to reduce the dough fermentation time, improve the properties of the dough and the crumb, in addition to the retention of aromas and moisture levels (Gerday et al., 2000). In bread production, in particular, microbial protease is used for modification of mixture containing high gluten content. Usually, blend containing high gluten will form a hard mixture. When protease is mixed in the blend, it will undergo partial hydrolysis and this will make the blend soft and easy to pull and knead. In addition to their use in breadmaking, enzymes are also used in biscuit manufacture, where the degradation of water-binding pentosans and partial hydrolysis of proteins improve product texture. A method has been patented that utilizes xylanase in reduced-fat or non-fat crackers to control their viscous and water-binding properties, thereby retaining a tender, non-brittle and shelf-stable structure (Beldman et al., 1996). For both of the above processes, the protease enzyme normally used is acidic protease from fungus. Protease is also frequently added to dough preparations. The enzyme has great impact on the quality of the bread and on dough rheology. Although the biochemical basis for these effects is not well known, it is probably due to effects on the gluten or gliadin fractions of the proteins. The protease used is a preparation from Aspergillus oryzae which has the effect of slackening the dough resulting in greater mobility and extensibility. Primo-Martín et al. (2006) has reported that proteolysis, which caused by proteases will lead to loss of protein structure, and a weaker, more open gluten network that improves crispness retention in bread crusts. However, excessive protease action causes stickiness or even soupiness in the dough (Poutanen, 1997).

Peptide Synthesis Since the first report of a pioneering work by Bergmann and his associates in 1937, on enzymatic synthesis of well-defined peptides using papain and α-chymotrypsin, proteases have emerged as powerful catalysts for the synthesis and modification of peptides (Isono and Nakajima, 2000). High concentrations of water-miscible organic solvents must be used in a biphasic system, however, to shift the chemical equilibrium away from hydrolysis toward synthesis. In the system, the enzymes are resided in the aqueous phase whereby the starting materials dissolved in the organic phase will diffuse into the water and undergo enzymecatalyzed peptide synthesis. Finally, the peptides formed will diffuse back into the organic phase from water. Enzymatic peptide synthesis offers several advantages over classical chemical methods, for instant, reactions can be performed stereospecifically and reactants do not require side-chain protection, increased solubility of non-polar substrates, or shifting

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thermodynamic equilibria to favor synthesis rather than hydrolysis (Gupta et al., 2002). These principles have been beautifully demonstrated by Cheetham (1994) in the large-scale production of aspartame. The stereospecific thermolysin catalyzed condensation between DLphenyalanine methyl ester (DL-PheOMe) and N-CBZ-L-aspartic acid gives exclusively the L, L-dipeptide product (N-CBZ-L-Asp-L-PheOMe). Though proteases has offered a number of important benefits, the applications of these enzymes in peptide synthesis has been hampered by some major limitations. The reliance on conventional multistep methodologies, the lack of knowledge regarding the specificity and properties of proteases in organic solvents and the strongly reduced activity of the enzyme under anhydrous conditions are among the mysteries that should be answered. Protein engineering holds great promise where the synthetic applications of these enzymes are concerned. For example, subtiligase, a double mutant in which the catalytic Ser221 is converted to Cys and Pro225 to Ala, displays a high catalytic constant and esterase to peptidase activity ratio and is an efficient peptide ligase in aqueous media (Abrahmsen et al., 1991). By using SDS-subtilisin as a catalyst for peptide bond formation in ethanol, Getun et al. (2001) have demonstrated that N-protected tri-peptides can be used directly as acyl donors, without the need to protect the peptide C-terminal. In addition, the enzyme is possible to synthesize tetra-peptides containing unprotected amino and carboxylic side groups in the P1- and P’1positions, again using ethanol as the solvent. A report by Xing et al. (2000) has indicated that zeolite immobilized α-chymotrypsin and thermolysin were active for peptide syntheses in organic media and still had catalytic activity to some extent after being reused five times.

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Varela, H., Ferrari, M. D., Belobrajdic, L., Vazquez A., and Loperena, M. L., 1997. Skin unhairing proteases of Bacillus subtilis: Production and partial characterization. Biotechnology Letter, 19: 755–758. Venugopal, M., and Saramma, A. V., 2006. Characterization of alkaline protease from Vibrio fluvialis strain VM10 isolated from a mangrove sediment sample and its application as a laundry detergent additive. Process Biochemistry, 41: 1239-1243. Visser, S., Robben, A. J., and Slagen, C. J., 1991. Specificity of a Cell-Envelope-Located Proteinase (PIII-type) from Lactococcus lactis subsp. cremoris AM1 in its action on Bovine β-Casein. Applied and Microbiology and Biotechnology, 35: 477-484. Xing, G. W., Li, X. W., Tian, G. L., and Ye, Y. H., 2000. Enzymatic peptide synthesis in organic solvent with different zeolites as immobilization matrixes. Tetrahedron, 56: 3517-3522. Yang, J., Teplyakov, A., and Quail, J. W., 1997. Crystal structure of the aspartic proteinase from Rhizomucor miehei at 2.15 Å resolution. Journal of Molecular Biology, 268: 449– 459.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 41-61

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 3

THERMOSTABLE LIPASES Thean Chor Leow, Fairolniza Mohd Shariff, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri ABSTRACT The stability of biocatalysts is an important criterion when dealing with bioprocesses at high temperature in order to sustain its operational activity throughout the processes. Much efforts has been focused on the screening of microorganisms harboring intrinsically stable biocatalysts. This chapter presents an overview of the issues involving screening, growth and production, purification and characterization of wild-type and recombinant enzymes with emphasis on thermostable lipases. High temperature, using olive oil as the sole carbon source, dictated the isolation of thermophilic lipolytic bacteria Geobacillus sp. strain T1 and Bacillus spp. strain 42 and strain L2. Tryptone and casamino acid were the best nitrogen sources, while corn oil and Tween 60 were the best substrates for the production of strain 42 lipase and L2 lipase, respectively. Molecular expression of thermophilic genes in mesophilic host not only reduced the exposure of recombinant enzymes to denaturing environment but facilitate protein purification and expression in bulk quantity in a shorter time. Theses lipases exhibited optimum temperature and pH of 70-80˚C and 7-9, respectively. These valuable properties create various potential industrial applications, particularly palm-based industry with respect to its high activity and substrate solubility at high temperature.

INTRODUCTION Since thermophilic microbial extracellular lipases are usually more thermostable than animal or plant lipases (Sugihara et al., 1991), they received much more attention for their potential application in the detergent, oil and fat, dairy and pharmaceutical industries (Gao et al., 2000). Furthermore, thermostable lipases are expected to play significant roles in industrial processing because running of bioprocesses at elevated temperatures lead to higher

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diffusion rates, increase in the solubility of lipids and other hydrophobic substrates in water and reduce the risk of contamination (Becker et al., 1997). In addition to higher thermostability, proteins from thermophiles often show better stability towards organic solvents and higher activity at elevated temperatures (Schmidt-Dannert et al., 1996). In addition, the melting point of fats varies significantly and can in some cases be as high as 50°C, but enzymatic catalysis on solid substrate is limited and therefore in those cases, it is difficult to use enzymes from mesophilic sources. However, thermophilic bacteria strains have optimum growth temperatures in the range of 65-70°C, which make lipases isolated from such strains good candidates for lipid modifications (Sigurgísladóttir et al., 1993). Therefore, screening and isolation of heat-stable lipase producers are important to fulfill industrial requirements.

SOURCES OF THERMOSTABLE LIPASES Microorganisms such as bacteria, fungi, yeast and actinomycetes have become possible sources of lipases (Sugihara et al., 1991). Microbial extracellular lipases are usually more thermostable than animal and plant lipases. In particular, lipases from thermophiles are expected to play significant roles in industrial processes since they are thermostable and resistant to chemical denaturation (Lee et al., 1999). In addition, the majority of lipases currently available are derived from mesophilic sources and display optimum activity at 35°C to 40°C (Sigurgísladóttir et al., 1993). Recently, microorganisms such as Bacillus sp. RSJ-1 (Sharma et al., 2001), Bacillus thermoleovorans ID-1 (Lee et al., 1999), Bacillus sp. THL027 (Dharmsthiti and Luchai, 1999), Bacillus thermocatenulatus (Schmidt-Dannert, 1994), Bacillus sp. strain A30-1 (Wang et al., 1995), Bacillus sp. Strain 398 (Kim et al., 1994), Bacillus spp. (Sugihara et al., 1991; Handelsman and Shoham, 1994; Nawani et al., 1998; Llarch et al., 1997; Becker et al., 1997), Pseudomonas cepacia (Sugihara et al., 1992), and Pseudomonas sp. (Kulkarni and Gadre, 1999), were reported as thermostable lipase producers. Besides that, moderate thermophiles can also be exploited to produce good sources of thermostable enzymes (Fakhreddine et al., 1998).

SCREENING AND ISOLATION In general, thermostable lipases are widely spread in nature from a variety of sources, for instance hot spring, soil, compost and effluent of palm oil mill. All these areas vary in relation to temperature, oil contamination, oil discharge or organic matter that may support the growth of heat-stable lipase producers. As lipases act not only on long chain triglycerides but also on those with medium and shorter carbon chains, various substrates could be used during preliminary screening processes. To date, tributyrin agar plate, Victoria Blue agar plate and Rhodamine B agar plate are normally used as screening plates with different sensitivity and selectivity. Although tributyrin agar plate confer more sensitivity but the hydrolysis of tributyrin can be catalysed by esterases, hence they may not show true lipase activity. Alternatively, a rather specific triolein agar plate with Victoria Blue as indicator which shows intense blue colour on pH reduction due to hydrolysis can also be used. On the other hand,

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triolein or olive oil agar plate with Rhodamine B as the indicator shows high selectivity but less sensitive as the orange fluorescence under UV, is hardly detected for low lipase activity.

GROWTH AND PRODUCTION OF WILD-TYPE LIPASE Growth factors either physical or nutritional can indeed make big impact on thermostable lipase production. Many studies to define the growth conditions that are optimum for thermostable lipases production were conducted. It is worth to note that production of thermostable lipases from thermophilic bacteria is normally less than that of mesophilic bacteria such as Pseudomonas sp., in which case lipase production of up to 8 U/mL was reported (Kulkarni and Gadre, 1999). To date, only limited studies were conducted on the production of thermostable lipases from thermophilic bacteria in shake-flask scale due to its low productivity. Therefore, lab-scale fermentors up to 15 L working volumes were normally preferred to promote better growth and overcome low level of production. Factors affecting production are described to gain insight into the growth and production of thermostable lipases.

Carbon Source Affecting Thermostable Lipase Production A thermophilic lipase from Bacillus thermocatenulatus showed fairly high lipase activity in the presence of 2.5 % (v/v) olive oil at 63°C after 44 h of cultivation at pH 6.5. The production was shown to be inducible with the addition of gum Arabic, where lipase activity increased by two times with 0.5 % gum Arabic and around 1.1 U/mL with 1 % gum Arabic. However, 0.5 % (w/v) Triton X-100 and 0.5 % (w/v) Tween-80 resulted in complete inhibition of lipase production (Schmidt-Dannert et al., 1994). On the other hand, the production of heat-stable lipase from Bacillus sp. strain H1 was around 0.4 U/mL in the presence of 1 % (w/v) Tween 80 but without olive oil after 36 h at 60°C (Handelsman and Shoham, 1994). A gram positive moderate thermophilic bacterium, LCB 39 produced significant lipolytic activity (31 U/L) when supplemented with 2 g/L glucose and 1 g/L Tween 80 to M9 mineral medium at 50°C under shaking condition but the lipase production (160 U/L) was significantly increased when glucose was omitted. The removal of glucose from MGT medium [2g/L glucose, 1g/L Tween 80 in mineral medium with the following composition (g/L): Na2HPO4 6; KH2PO4 3; NaCl 0.5; NH4Cl 1)] resulting in the increase of lipase activity suggested an inhibitory effect of glucose such as catabolite repression. However, the lipase production was further increased to 300 U/L by increasing the Tween 80 concentration up to 8 g/L (Fakhreddine et al., 1998). Dharmsthiti and Luchai (1999) reported a low cost medium, MGRS [0.1 % glucose (w/v) and 1 % rice bran oil (v/v) as carbon sources] supplemented with 0.5 % (w/v) rice bran for Bacillus THL027 to produce a lipase activity of 7.8 U/mL as compared to the more expensive nutrient broth. Kumar et al. (2005) reported the highest lipase production of 1.16 U/mL by B. coagulans BTS-3 in shake flask scale when refined mustard oil was used as carbon source at 55°C and pH 8.5 after 48 h. A highly stable alkaline lipase was reported from a newly isolated thermophilic Bacillus sp. strain A30-1 from a hot spring area in Yellowstone National Park (Wang et al., 1995). It

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produced extracellular lipolytic enzyme and was able to utilize corn oil as substrate. The lipase production was nearly 3.5 U/mL in 1 L fermentor with 500 mL of working volume (pH 9.0) with 1 % (v/v) corn oil, 0.1 % (v/v) Tween 80 and 0.1 % (w/v) yeast extract at 60°C. Becker et al. (1997) reported a new isolate Bacillus sp. strain IHI-91 that grew optimally at 65°C and pH 6-6.5. The duration of a typical batch fermentation with 1 % (v/v) olive oil in defined salt medium (1.6 L) at 65°C exhibited a maximum lipase activity of 0.3 U/mL after 78 h of fermentation. Another thermopilic microorganism, B. thermoleovorans ID-1, isolated from a hot spring in Indonesia, showed extracellular lipase activity and high growth rates on lipid substrates at elevated temperatures (Lee et al., 1999). The lipase activity reached the maximum value of 512 U/L during the late exponential phase with olive oil (1.5 %, v/v) as the sole carbon source in a 2 L bioreactor with a working volume of 1.5 L. Thermophilic Bacillus sp. RSJ-1 was cultivated in 20 L laboratory bioreactor with 15 L lipase production medium (pH 9.0) containing 0.75 % (v/v) cottonseed oil and 0.5 % Tween-80 (v/v) as working volume. A lipase activity of around 10.2 U/mL was detected after 10 h of cultivation time at 50°C (Sharma et al., 2001). Besides the above mentioned thermostable lipase producers, mesophilic sources, Pseudomonas spp. indeed may serve as a good candidate for heat-stable lipase producers. Gilbert et al. (1993) used response surface methodology to optimize the lipase production of Pseudomonas aeruginosa EF2 growing in continuous culture showed that maximal lipase activity (39 U/mg cells) occurred during growth under Tween limitation at low dilution rate (0.04 h-1) at pH 6.5 and 35.5°C. In addition, a hyper-thermostable alkaline lipase from Pseudomonas sp. was able to produce a specific activity of 24 U/mg when growth in medium containing glucose (10 g/L) and coconut oil (20 mL in 1 L) at 50°C, pH 7 under shaking condition (250 rpm) for 24 h (Rathi et al., 2000). A novel alkaline, thermostable, proteasefree lipase from Pseudomonas sp. was also reported by Kulkarni and Gadre (1999). In shake flask culture, addition of groundnut oil (3 g/L) towards the end of the log phase increased the lipase activity from 4 U/mL to 8 U/mL.

Nitrogen Source Affecting Thermostable Lipase Production Several nitrogeneous compounds such as ammonium salts, yeast extract, peptone and casamino acids are widely used as nitrogen sources for the above mentioned thermostable lipases production regardless shake-flask or fermentor scale. Bacillus thermoleovorans ID-1 and Bacillus sp. RSJ-1 preferred a combination of complex nitrogen sources of 6 g/L tryptone + 2 g/L yeast extract and 0.75 % peptone + 0.75 % yeast extract in their fermentation medium, respectively (Sharma et al., 2001; Lee et al., 1999). In agreement to Bacillus RSJ-1, a combination of 0.5 % (w/v) peptone and 0.5 % yeast extract (w/v) synergically promoted higher lipase production from B. coagulans BTS-3 (Kumar et al., 2005). In addition, lipase production medium containing of 0.4 % (w/v) casimino acids or 0.1 % (w/v) yeast extract could also be used to cultivate Bacillus sp. H1 and Bacillus sp. A30-1 (Wang et al., 1995; Handelsman and Shoman, 1994). Besides utilization of complex nitrogen sources, some strains such as Bacillus sp. IHI-91 and Bacillus sp. THL027 used 1 g/L and 0.1 % (w/v) of ammonium sulphate, respectively as nitrogen source for lipase production (Dharmsthiti and Luchai, 1999; Becker et al., 1997). However, additional nitrogen source of 0.5 % (w/v) rice

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bran was also used together with ammonium salt by Bacillus sp. THL027. On the other hand, Rathi et al. (2000) reported a lipase production medium containing of a combination of inorganic sources 5 g/L ammonium chloride and 3 g/L ammonium phosphate for Pseudomonas sp.

GROWTH AND PRODUCTION OF RECOMBINANT LIPASE Microbes are extremely good at producing an amazing array of compounds; however, as the compounds are mainly for their own use, the quantities are small. Therefore, recombinant DNA manipulation can be used to increase the level of production. Many lipase genes have been cloned and expressed over the past few years. Thermostable lipase from Bacillus stearothermophilus L1 was not purified from wildtype bacteria due to its low level of production as well as its strong interaction with other proteins and lipid materials. Therefore, a structural gene of L1 mature lipase was expressed in pET22b(+) and its expression was induced with Isopropyl β-D-thiogalactopyranoside (IPTG) at 37°C in Escherichia coli BL21(De3) cells. An expression level of 132,000 U/mL was obtained through recombinant DNA technology as compared to less than 200 U/L for wildtype lipase (Kim et al., 2000). However, Cho et al. (2000) reported that the recombinant lipase activity was approximately 1.4-fold greater than under the native promoter when expressed in the same vector as L1 lipase. Heat-stable lipase of B. stearothermophilus P1 was cloned into the expression vector pQE-60 prior to use to transform Escherichia coli M15 [pREP4]. Overexpression of the cloned lipase P1 induced by IPTG addition resulted in a high expression of soluble lipase activity of 212 x 103 U/L as compared with 8.1 U/L using B. stearothermophilus P1 (Sinchaikul et al., 2001). A lipase-producing thermophilic strain TW1, assigned as Geobacillus sp. TW1 based on its 16S rRNA analysis sequence, was isolated from a hot spring in China. Its open reading frame encoding 417 amino acids was expressed in E. coli as a glutathione S-transferase fusion protein with E. coli BL21(De3) as a host (Li and Zhang, 2005) . BTL2 lipase from B. thermocatenulatus was expressed in pCYTEXP1 expression vector downstream of the temperature-inducible λ promoter PL. The expression level encountered was around 9000 and 7000 U/g cells for BTL2 lipase with and without signal peptide, respectively. However, significantly improved in expression was observed when OmpA protein (for translocation to the periplasm) was fused to the BTL2 gene encoding the mature lipase (Rua et al., 1998).

PURIFICATION AND CHARACTERIZATION OF THERMOSTABLE LIPASE Wild-type Lipase A thermostable lipase from Bacillus thermoleovorans ID-1 was purified to homogeneity through ammonium sulphate fractionation, DEAE-Sephacel and Sephacryl S-200 (Lee et al., 1999). A single band on SDS-PAGE was estimated to be about 34 kDa. The highest activity

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was obtained at 75°C and stable up to 50°C, while retaining 73 and 50 % of its original activity at 60 and 70°C, respectively. The lipase showed the highest activity towards pnitrophenyl caproate (C6 acyl group) but preferred tricaprylin (C8) among the triacylglycerols tested. The catalytic activity increased to about 120% in the presence of Ca2+ and Zn2+. SDS and PMSF at 1 mM inhibited wild-type lipase from B. thermoleovorans ID-1. THL027 lipase was purified by 2.6-fold with 27% yield and a specific activity of 4.5 U/mg by a single step of Sephadex G-100 gel filtration prior to ultrafiltration through 10 kDa cut-off membrane (Dharmsthiti and Luchai, 1999). The molecular mass of THL027 lipase determined by SDS-PAGE to be 69 kDa. It was highly active at 70°C and pH 7. With respect to fatty acid specificity, the enzyme hydrolysed triglycerides containing short- to mediumchain saturated fatty acids (C4-C12) faster than it did for saturated long-chain fatty acids (C14C18). Tricaprylin (C8:0) and olive oil being the best triglyceride and natural oil, respectively. B. thermocatenulatus culture concentrate was stored for 2 days at 4°C. The newly formed calcium soap was separated by filtration of the concentrate prior to extraction with hexane. During phase separation, a small intermediate phase containing lipase was further purified with methanol precipitation and ion-exchange chromatography Q-Sepharose column. The final recovery of 11 % and specific activity of 29 U/mg were obtained. The purified lipase with molecular weight of 16 kDa showed an optimum activity at 70 and 65% with olive oil and p-NPP as substrates respectively. It was stable up to 40°C in phosphate buffer at pH 7.5 for 0.5 h. Linseed oil and soybean oil were the preferred substrates compared to fish oil and triglyceride triolein. Addition of 1 % (w/v) detergent octylglucoside, CHAPS and taurocholate activated the lipase activity immediately but became inactivated in the presence of Tween 20 and 80 under the same concentration. However, Brij 35, Triton X-100 and Lubrol PX inactivated the lipase slightly in the incubation mixture. The lipase forms very large aggregates (>750 kDa) as observed on native PAGE. Therefore, activity display will not be the true lipase value due to aggregate formation, as only lipase molecules at the surface of the aggregates are available for the substrate (Schmidt-Dannert et al., 1994). An extracellular, thermostable, alkaline lipase was partially purified from a thermophilic Bacillus strain J33 (Nawani et al., 1998). It was partially purified 8-fold by ammonium sulphate and 10-fold by gel filtration with final specific activity of 197 U/mg. This native enzyme has an apparent relative molecular mass of 45 kDa. The optimum temperature was 60°C at pH 8, stable up to 60°C and has a half-life of 30 min at 70°C. The lipase was stable in 90 % (v/v) hexane or benzene mixtures in water. Metal ions Mg2+, Na+ and Li+ increased lipase activity by 63%, 63% and 23% respectively. Strong inhibition effect was, however, exhibited by Hg2+ and Cd2+. EDTA and PMSF at 10 mM severely inhibited J33 lipase activity. An extracellular alkaline lipase from a new thermophilic Bacillus sp. RSJ-1 was purified to homogeneity by ultrafiltration, followed by ammonium sulphate precipitation, dialysis, QSepharose ion exchange chromatography and Sephacryl S-200 SF gel filtration chromatography (Sharma et al., 2001). The final yield of 19.7 % and purification fold of 201 were achieved by the purification protocols. The molecular mass of RSJ-1 lipase was determined to be 37 kDa by SDS-PAGE, with an optimum temperature and pH of around 50°C and pH 8, respectively. The enzyme activity was increased in the presence of 1 mM metal ions Na+, Mg2+ and Ba2+ by 12, 2, and 2 %, respectively. However, K+, Co2+ and Zn2+ strongly inhibited its activity. Oxidizing agents (ammonium persulphate, potassium iodide)

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and reducing agents (ascorbic acid, 2-mercaptoethanol) moderately inhibited lipase activity compared to chelating agents (sodium citrate, EDTA), where no significant lost of activity was detected. It was stable in various surfactants and commercial detergents and hence has great potential in detergent formulation. A thermophilic isolate, B. coagulans BTS-3 was able to produce extracellular alkaline lipase (Kumar et al., 2005). The enzyme was purified 40-fold to homogeneity by ammonium sulphate precipitation and DEAE-Sepharose column chromatography. The molecular weight was 31 kDa on SDS-PAGE. The enzyme exhibited an optimum activity at 55°C and pH 8.5. The half-life at 55°C was around 2 h. Al3+, Co2+, Mn2+ and Zn2+ were found to inhibit the activity, while K+, Fe3+, Hg2+ and Mg2+ enhanced the enzyme activity. However, Na+ has no effect on its activity. The BTS-3 lipase has a preference towards long chain synthetic substrates, especially 4-nitrophenyl palmitate. Handelsman and Shoham (1994) reported a partially purified H1 lipase by acetone precipitation, anion exchange chromatography and gel filtration with specific activity of 0.95 U/mg. The enzyme has a molecular weight of 20,000 Da when calibrated with standard protein by gel filtration. The H1 lipase was most active at pH 7.0 at 70°C with a half-life of 50 h at 60°C. The metal ions Ca2+, Na+, K+, Mg2+, Co2+ and Ni2+ showed a slight enhancing effect on the activity, but Ba2+ enhanced the activity by 32 %, while Hg2+, Al3+ and Fe2+ inhibited the activity. The lipase from Bacillus A30-1 could be efficiently recovered by ultrafiltration of cellfree culture supernatant followed by ammonium sulphate precipitation and then subjected to a 8 % polyacrylamide preparative gel electrophoresis performed on a vertical slab gel apparatus to recover the most active and highest specific activity. The partially purified lipase had an optimum temperature of 60°C, at an optimum pH of 9.5. It retained 100% of its original activity after being heated at 75°C for half an hour. The half-life was around 8 h at 75°C. It preferred triglycerides trilaurin as well as natural fats and oils as substrates (Wang et al., 1995). A hyper-thermostable lipase, alkaline lipase from mesophilic Pseudomonas sp. was partially purified by ultrafiltration (10 kDa cut-off) and thermal precipitation for 1 h at pH 11. It was 6.8-fold with a recovery of 71 %. The optimum activity was at 90°C and pH 11. The enzyme retained 100% of its original activity upon treatment for 8 h at 90°C with a half-life of more than 13 h. This lipase from Pseudomonas sp. catalysed the hydrolysis of mustard and linseed oil most efficiently followed by neem, castor, groundnut and coconut oils (Rathi et al., 2000). A number of extracellular microbial thermostable lipases was obtained from various sources, especially from Bacillus spp. and Pseudomonas spp. Many researchers have studied the properties of crude and purified microbial lipases. These properties are critical in the final evaluation on the suitability of the enzymes for industrial applications. Some such properties of thermostable lipases are presented in Table 1. As can be seen, the optimal activity for Bacillus spp. and Pseudomonas spp. are obtained at pH ranging from pH 7.0 to 9.5 and 9.6 to 11.0 respectively and temperatures ranging from 50 to 90 °C for both genera. Since most of the thermostable lipases reported so far come from Bacillus spp. and Pseudomonas spp. hence it is not only the thermophilic bacteria but also the mesophiles such as Pseudomonas spp. can become possible sources of thermostable lipases.

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Recombinant Lipase Recombinant L1 lipase of Bacillus stearothermophilus was overexpressed in Escherichia coli harboring recombinant plasmid pET22b(+). The L1 lipase was efficiently purified to homogeneity with CM-Sepharose (pH 6.0) and DEAE-Sepharose (pH 8.8) column chromatographies with a recovery of 70.1 and 62.2 %, respectively. The specific activity of purified lipase was 1700 U/mg when olive oil was used as substrate. The recombinant L1 lipase showed an optimum temperature of 68°C at pH 8. It was stable up to 55°C for 30 min of incubation time. Its thermal stability increased by 8-10 degrees in the presence of calcium ions (Kim et al., 2000). Table 1: Properties of thermostable lipases from different sources.

Organism

Crude lipase: Bacilus sp. strain A30-1a Bacillus sp. Strain J33b Pseudomonas sp.c Pseudomonas sp.d Purified lipase: Bacillus sp. strain 398e Bacillus thermocatenulatusf Bacillus stearothermophilus L1g Bacillus sp. THL027h Bacillus sp. RSJ-1i Bacillus thermoleovorans ID-1j

Enzyme properties Optimal Optimal temperature(°C ) pH

Temperature stability

60 60 65 90

9.5 8 9.6 11

T1/2 ~ 8 h at 75 °C T1/2 ~ 30 min at 70 °C T1/2 ~ 50 min at 75 °C T1/2 ~ 13 h at 90 °C

65 60-70 65 70 50 60

9 7.5-8.0 9.5 7 8-9 9

T1/2 ~ 30 min at 65 °C T1/2 ~ 30 min at 60 °C T1/2 ~ 30 min at 63 °C T1/2 ~ 30 min at 75 °C T1/2 ~ 30 min at 70 °C

a: Wang et al., 1995 b: Nawani et al., 1998 c: Kulkarni and Gadre, 1999 d: Rathi et al., 2000 e: Kim et al., 1994 f: Schmidt-Dannert et al., 1994 g: Kim et al., 1998 h: Dharmsthiti and Luchai, 1999 i: Sharma et al., 2001 j: Lee et al., 2001

Escherichia coli BL321 was transformed with the expression plasmid pCYTEXP1 carrying BTL2 gene from B. thermocatenulatus under the control of the strong temperatureinducible λPL promoter (Rúa et al., 1998). The mature lipase was purified to homogeneity by a two-step purification protocol of hydrophobic interaction chromatography and gel filtration chromatography. The final yield and purification fold were around 32% and 125 respectively at specific activity of 54,887 U/mg. However, the lipase has the tendency to form active molecular aggregates. This recombinant BTL2 lipase exhibited the maximum activity at pH 8.0 and pH 9.5 with tributyrin and triolein, respectively, and an optimum temperature of 55°C

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and 75°C for the hydrolysis of tributyrin and triolein respectively. Although the optimum temperature was high, it was only stable up to 50°C when incubated for 30 min at pH 9.0. Tricaprylin (C8) was the preferred substrate compared to other triglycerides tested, regardless of the assay at pH of 7.5 or 8.5. The addition of Cholate, CHAPS, Triton X-100 or OGP resulted in increased activity ranging from 40% (CHAPS and OGP) to 80% (Triton X-100 and cholate). However, SDS, Lubrol PX, Tween 20 or 80 immediately deactivated the enzyme. In addition, the addition of 30% (v/v) methanol, 30% (v/v) 2-propanol or 30% (v/v) acetone had a slight inhibitory effect on the BTL2 lipase activity. P1 lipase was overexpressed in prokaryotic system harboring recombinant plasmid pQE60 (Sinchaikul et al., 2001). The recombinant lipase was successfully purified through a single chromatography step using a strong anion exchanger Q-HyperD10. The recombinant lipase exhibited specific activity of 811 U/mg after being purified by 18-fold with a recovery of 71%. The molecular mass of the lipase was 43 kDa by SDS-PAGE and mass spectrometry. The purified lipase had an optimum pH of 8.5 and showed maximum activity at 55°C. The P1 lipase was stable ranging from 30 to 65°C. The highest activity was found with p-nitrophenyl ester-caprate as synthetic substrate and tricaprylin as the triacylglycerol. Zn2+ and Fe2+ significantly inhibited its activity. It was stable in the presence of 0.1% CHAPS, Triton X-100 for 1 h. Metal ions Ca2+, Mg2+, and Na+ stabilized its lipase activity, but Cu2+, Cs2+, K+ slightly inhibited P1 lipase activity. PMSF, 1-Dodecanesulfonyl chloride and 1Hexadecanesulfonyl chloride at 10 mM strongly inhibited P1 lipase.

OUR WORK Screening of Lipolytic Enzymes We have conducted an extensive screening programme for thermophilic microbes, mainly bacteria, as possible sources of microbial lipases for industrial application. Sampling sites include hot springs, effluent from palm oil mills, palm oil branches and rubbish dump sites with temperatures up to 91°C for hot springs and 48°C for the latter (Figure 1). Enrichment medium with the sole carbon source from olive oil was used to enrich these heat stable lipase producers under shaking condition. Screening agar plates such as tributyrin, Victoria Blue (Figure 2) and Rhodamine B agar plates were used in the preliminary selection. Three thermostable lipase producers namely Geobacillus sp. strain T1, Bacillus sp. strain 42 and Bacillus sp. strain L2 were successfully isolated.

Growth and Production of Thermostable Lipases Wild-type Lipase A thermophilic microorganism, Bacillus sp. strain L2 was isolated from a hot spring in Perak, Malaysia. A number of factors affecting the production of its extracellular lipase were investigated. The optimum lipase production was obtained at 70°C, in pH 7 medium, 150 rpm of agitation rate, 1 % of starting inoculum size for 28 h incubation time. In addition, casamino

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acids, trehalose, Ca2+ and Tween 60 were found to be effective nutrition for lipase production. A final production of 0.158 U/mL was obtained after optimization studies.

A) Hot spring in Slim River, Perak

B) Palm oil mill effluent at Semenyih, Selangor

C) Palm fruit branches near palm oil factory at Semenyih, Selangor Figure 1: Sampling sites for thermostable lipase producers

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Productions of thermostable lipases derived from thermophilic bacteria are usually low, as in the case of Bacillus sp. strain 42. Heat denaturation after long exposure at high temperature could be the main reason. Optimization of lipase production is paramount to yield sufficient enzyme for downstream laboratory processing, particularly for protein purification prior to enzyme characterization and protein crystallization. In order to increase the production of strain 42 lipase, carefully designed experimental parameters involving various carbon sources, nitrogen sources, metal ions and natural oils are taken into consideration (Eltaweel et al., 2005).

Figure 2: Victoria Blue agar plate showing putative lipase producer. Geobacillus sp. strain T1 formed intense blue color around colonies with Bacillus sp. strain F1 as negative control.

The effect of additional carbon sources in the basal medium for lipase production by Bacillus sp. strain 42 is as shown in Table 2. Sorbitol, fructose and arabinose did not improve lipase production with respect to the basal medium. However, addition of mannitol and lactose encountered 9% and 23% reduction in lipase production, respectively. Addition of starch, rhamnose, melibiose and myo-inositole decreased the lipase production about 32 to 46% compared to the basal medium, while 58 to 60% reduction was observed with the addition of raffinose and ethanol. Lipase production was much lower with trehalose and galactose. The other carbon sources tested such as maltose, mannose, sucrose and glucose completely inhibited lipase production. The effects of various nitrogen sources on lipase production by Bacillus sp. strain 42 was shown in Figure 3. The addition of tryptone increased lipase production about 45% compared to the basal medium. In addition, soytone, corn steep liquor, proteose peptone, beef extract and peptone also enhanced lipase production about 35, 23, 17, 10, 6%, respectively, as compared to basal medium. However, addition of yeast extract decreased the lipase production to 41%, while casein, casamino acid and mollases completely inhibited the lipase production. The inhibition phenomena could be due to the inability of the bacterium to hydrolyse them, or that the amino acids released were toxic to the bacterium. Another possibility is that the amino acids released

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into the medium could have denatured the lipase formed. Inorganic nitrogen sources used in this study such as sodium nitrate and di-ammonium orthophosphate drastically reduced lipase production to 86% and 95%, respectively, while other inorganic sources totally inhibited lipase production (Eltaweel et al., 2005). Table 2: Effect of carbon sources as additives (1%, w/v) to the basal medium on lipase production. Carbon source Basal medium Arabinose Fructose Sorbitol Mannitol Lactose Myo-inositol Melibiose Rhamnose Starch Ethanol Raffinose Trehalose Galactose Glucose Sucrose Mannose Maltose

Relative activity (%) 100 99 98 98 91 77 68 57 56 54 42 40 25 22 0 0 0 0

Tryptone was identified earlier as the best nitrogen source (Figure 3) for Bacillus sp. strain 42. In this study, each of the nitrogen sources (soytone, peptone, beef extract, proteose peptone, corn steep liquor and yeast extract), which supported lipase production individually, were added at 0.4% each to tryptone production medium. Tryptone and yeast extract together was found to be the best for lipase production. This combination increased the lipase yield to 1.8 fold as compared to medium containing only tryptone. The percentage of tryptone and yeast extract was then optimised. Table 3 shows that a combination of 0.6% of tryptone and 0.2% yeast extract was the best for enzyme production. Tryptone seemed to play an important role in lipase synthesis, while yeast extract supported bacterial growth, resulting in higher lipase production (Eltaweel et al., 2005). Metal ions are important for the production of lipase. In this study, the basal medium used as control contained a mixture of metal ions Mg2+, Ca2+ and Fe3+. When only individual ions are utilised, the medium could only support lower lipase production of which Fe3+ was the best at 84% of the basal production medium (Figure 4). A combination of two metal ions added to the culture medium, e.g. Ca2+ + Fe3+, Ca2+ + Mg2+, and Mg2+ + Fe3+ gave lower production of 42 lipase. Therefore, the basal medium was formulated with all three ions to ensure good lipase production by Bacillus sp. strain 42. In this study, addition of other heavy

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metal ions to the basal medium such as Co2+, Cu2+, Mg2+ and Zn2+ were found to support low lipase production in 24 h.

Figure 3: Effect of nitrogen sources as additives (0.8%, w/v) to the basal medium (without nitrogen source) on lipase production.

Natural oils when utilised as substrates had different effects on lipase production. A study on the production of 42 lipase by Bacillus sp. strain 42 showed that olive oil was the best substrate (Figure 5). Corn oil and crude palm oil supported considerable amount of lipase production up to 88% and 77% of olive oil in basal medium, respectively. As shown, most of the substrates tested supported lipase production in the range of 44 to 88% of basal medium (olive oil). However, tributyrin and crude palm kernel stearin totally inhibited the lipase production. Table 3: Effect of different percentage of tryptone and yeast extract combination on lipase production by Bacillus sp. strain 42 Tryptone (%) 0.1 0.2 0.3 0.4 0.5 0.6 0.7 (Source: Eltaweel et al., 2005)

Yeast extract (%) 0.7 0.6 0.5 0.4 0.3 0.2 0.1

Lipase activity (U/ml) 0.097 0.099 0.110 0.132 0.165 0.174 0.119

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Figure 4: Effect of metal ions used individually and in combination in the basal medium: 0.02% Ca2+, 0.04% Fe3+, 0.01% Mg2+, 0.02% Ca2+ + 0.04% Fe3+, 0.02% Ca2+ + 0.01% Mg2+ and 0.01% Mg2+ + 0.04% Fe3+ [Fe3+ prepared as 1% (w/v) stock solution].

Figure 5: Effect of substrates on lipase production by Bacillus sp. strain 42. Olive oil in the basal medium was substituted with various substrates.

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Recombinant Lipase For lipase production by Geobacillus sp. strain T1, the bacterium was grown aerobically in basal mineral medium composed of (g/L) : NaNO3 : 7; K2HPO4 : 2; KH2PO4 : 1; KCl : 0.1; MgSO4.7H2O : 0.5; CaCl2 : 0.01; FeSO4.7H2O : 0.012; yeast extract : 1 in which 0.01 % trace elements and 2 % (v/v) olive oil were supplemented. Low production level of 0.15 U/mL was detected after 24 h of incubation time. Therefore, molecular cloning offers an alternative route of production economically in a shorter time. An open reading frame of thermostable T1 lipase gene was initially cloned into pBAD TOPO TA vector through PCR cloning method. The recombinant clone obtained not only formed clearing zones but also intense blue color and orange fluorescence around positive clones on tributyrin, Victoria Blue and Rhadamine B agar plates respectively (Figure 6). A comparative expression was conducted in pBAD, pRSET, pET, and pGEX which under the control of araBAD, T7, T7lac and tac, respectively (Leow et al., 2004). The optimum expression level for pBAD/T1, pRSET/T1, pET/T1 and pGEX/T1 was 1.941, 2.366, 0.644 and 5.625 U/mL respectively (Table 4). Further optimization was performed with recombinant clone Escherichia coli (pGEX/T1) by varying the inducer IPTG concentration. Low inducer concentrations ranging from 0.025 to 0.1 mM were more effective in regulating the transcription and translation processes of T1 lipase gene. The pGEX system was not a tight expression system since a significant level of basal transcription (6.872 U/mL) was detected in uninduced culture after 8 h of induction time. An inducer concentration as low as 0.05 mM was sufficient to induce the recombinant culture, with optimum expression level (11.708 U/mL) detected upon 8 h of post-induction time. Indeed, a higher intensity for the expressed band can be seen on lane 3 of Figure 7. The molecular mass of fusion lipase (signal peptide) and GST tag was around 63 kDa and 26 kDa, respectively. There was a two times enhancement in expression with inducer as low as 0.05 mM after 8 h of induction time at 37˚C. The highest production level obtained was around 78 folds as compared to wild-type Geobacillus sp. strain T1. Extracellular production of T1 lipase was by coexpression of pJL3 vector encoding bacteriocin release protein (BRP) in prokaryotic system. Secretory expression was optimized by considering several parameters, including host strains, inducer (IPTG) concentration, media, induction at A600nm, temperature and time of induction (Rahman et al, 2005). Among the host strains tested, E. coli Origami B excreted out 18.100 U/ml of lipase activity into culture medium when induced with 50 µM of IPTG for 12 h. The E. coli Origami B harboring recombinant plasmid pGEX/T1S and pJL3 vector was chosen for further study. IPTG at 0.05 mM, YT medium, induction at A600nm of 1.25, 30°C and 32 h of induction time was the best condition for T1 lipase secretion with Origami B as host (Rahman et al., 2005). Figure 8 showed an expression profile of E. coli Origami B harboring recombinant plasmid pGEX/T1S and pJL3 vectors intra and extracellularly. Initially, the intracellular level expression was higher compared to extracellular expression. This was due to the quasi-lysis effect generated by bacteriocin released protein to activate phospholipase A, forming weak permeable zone with low amount of inducer. As a consequence, the longer induction has promoted higher level of active protein extracellularly at 32 h of cultivation time. However, a moderate induction prevented the lysis of producer cells and maintained the viability of the culture, making the system suitable for large-scale protein production in a continuous culture.

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pBAD/T1

pBAD/lacZ A) Tributyrin agar plate

pBAD/T1

pBAD/lacZ

B) Victoria Blue agar plate

pBAD/T1

pBAD/lacZ C) Rhodamine B agar plate

Figure 6: Screening plates showing positive recombinant clone. A) Tributyrin agar plate; B) Victoria Blue agar plate; C) Rhodamine B agar plate

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Table 4: A summary of optimizing expression of T1 lipase gene with signal peptide. Expression system pBAD/T1 pRSET/T1 pET/T1 pGEX/T1

Promoter araBAD T7 T7lac tac

Host strain Top 10 BL21(De3) BL21(De3)pLysS BL21(De3) BL21(De3)pLysS BL21(De3) BL21(De3)pLysS BL21(De3)pLysS

Lipase activity (U/mL) 1.941 (36 h) 1.960 (20h) 2.366 (12 h) 0.231 (8 h) 0.644 (8 h) 4.077 (12 h) 5.625 (8 h) 11.708 (0.05 mM)

Note: Crude cell lysates were prepared with a sonicator. The soluble fractions were assayed colorimetrically with olive oil as substrate. One unit of lipase activity was defined as 1µmole of liberated fatty acid per minute the under the assay conditions. The cultures were induced at an A600nm ~ 0.5 with 0.02 % of L-arabinose (pBAD) and 1 mM of IPTG (pRSET , pET, pGEX) at 37 °C. The optimal induction time and inducer concentration are in bracket. (Source: Leow et al., 2004)

Figure 7: SDS-PAGE (12%) of expressed T1 lipase. M: standard protein markers were β-galactosidase (116 kDa), bovine serum albumin (66.2 kDa), ovalbumin (45 kDa), lactate dehydrogenase (35 kDa), restriction endonuclease Bsp 98l (25 kDa), and β-lactoglobulin (18.4 kDa), without IPTG (lane 1), 0.025 mM (lane 2), 0.050 mM (lane 3), 0.100 mM (lane 4), 0.500 mM (lane 5), 1 mM (lane 6), 1.5 mM (lane 7), 2 mM (lane 8) and GST (lane 9). Arrows indicate GST fusion protein and GST. (Source: Leow et al., 2004)

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Figure 8: Effect of induction times on T1 lipase production by E. coli Origami B harboring recombinant plasmid pGEX/T1S and pJL3 vectors. The cultures were analysed intracellularly ( ) and extracellularly ( ) after induced with 50 µM of IPTG at A600nm of 1.25 and incubated at 30˚C up to 40 h. (Source: Rahman et al., 2005)

Purification and Characterization The findings demonstrate that molecular biotechnology offers better alternative route towards production or purification to overcome low lipase production of wild-type bacteria. Cloning and expression in pQE-30 UA vector conferred single step protein purification economically through Ni-NTA affinity chromatography technique. The 42 lipase was highly active at 70°C and pH 8.0. It was stable in the presence of Ca2+ and 0.1% (v/v) Tween 60 and 80 but moderately inhibited by Tween 20, 40 and 85. The 42 lipase was an organic solvent tolerant lipase, stable in various solvents tested. The purified 42 lipase preferred olive oil and tricaprylin for catalysis. However, it showed random substrate specificity when evaluated with TLC. Besides that another thermostable lipase was partially characterized with respect to its crude enzyme. The crude recombinant T1 lipase showed an optimum temperature of 65°C when olive oil was used as substrate and stable up to 65°C for 30 min. The optimum pH of crude fusion T1 lipase was around 9. It was stable over a wide pH range of 6 to 11 but the stability decreased drastically above pH 11 (Leow et al., 2004). T1 mature lipase was purified through affinity chromatography. The purified T1 mature lipase displayed an optimum temperature of 70oC and pH of 9. The half-live at 65˚C was about 5 h 15 min. The wild-type L2 lipase from Bacillus sp. L2 displayed an optimum temperature and pH of 80˚C and pH 7 respectively. It was highly stable for 2 h at 80˚C. Metal ions Fe2+ and Ca2+ promoted L2 lipase activity while Cu2+ inhibited it. However, recombinant L2 lipase exhibited lower temperature activity and stability at 70˚C and a half-life of 30 min at 60˚C, suggesting that the recombinant L2 lipase might need post translational modification such as glycocylation for protein solubility and stability.

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Some characteristics of the above mentioned thermostable lipases are listed in Table 5. The crude and purified lipases are highly active at pH 7 to 9. The optimum temperatures for crude lipases and purified lipases were 70-80°C and 65-70°C respectively. These properties create potential industrial applications of these lipases. Table 5: Properties of thermostable lipases from different sources.

Organism

Purified lipase: T1 lipase 42 lipase L2 lipase (wild-type) Crude lipase: T1 fusion lipase (recombinant) L2 lipase (recombinant)

Enzyme properties Optimal Optimal temperature ( °C ) pH

Temperature stability

70 70 80

9 8 7

T1/2 ~ 5 h 15 min at 65 °C T1/2 ~ 2 h at 65 °C T1/2 ~ 2 h 40 min at 80 °C

65 70

9 7

T1/2 ~ 30 min at 68 °C T1/2 ~ 30 min at 60 °C

REFERENCES Becker, P., Reesh, I. A., Markossian, S., Antranikian, G., and Märkl, H., 1997. Determination of the kinetic parameters during continuous cultivation of the lipase-producing thermophile Bacillus sp. IHI-91 on olive oil. Applied Microbiology and Biotechnology, 48: 184-190. Cho, A. R., Yoo, S. K., and Kim, E. J., 2000. Cloning, sequencing and expression in Escherichia coli of a thermophilic lipase from Bacillus thermoleovorans ID-1. FEMS Microbiology Letter, 179: 235-238. Dharmsthiti, S., and Luchai, S., 1999. Production, purification and characterization of thermophilic lipase from Bacillus sp. THL027. FEMS Microbiology Letters, 179: 241246. Eltaweel, M. A., Rahman, R. N. Z. R. A., Salleh, A. B., and Basri, M., 2005. An organic solvent-stable lipase from Bacillus sp. strain 42. Annals of Microbiology, 55 (3): 187-192. Fakhreddine, L., Kademi, A., Abdelkader, N. A., and Baratti, J. C., 1998. Microbial growth and lipolytic activities of moderate thermophilic bacterial strains. Biotechnology letters, 20 (9): 879-883. Gao, X. G., Cao, S. G., and Zhang, K. C., 2000. Production, properties and application to nonaqueous enzymatic catalysis of lipase from a newly isolated Pseudomonas strain. Enzyme and Microbial Technology, 27: 74-82. Gilbert, E. J., 1993. Pseudomonas lipases: Biochemical properties and molecular cloning. Enzyme and Microbial Technology, 15: 634-645. Haki, G. D., and Rakshit, S. K., 2003. Developments in industrially important thermostable enzymes: a review. Bioresource Technology, 89: 17-34.

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Handelsman, T., and Shoham, Y., 1994. Production and characterization of an extracellular thermostable lipase from a thermophilic Bacillus sp. Journal of General Applied Microbiology, 40: 435-443. Kim, H. K., Park, S. Y., Lee, J. K., and Oh, T. K., 1998. Gene cloning and characterization of thermostable lipase from Bacillus stearothermophilus L1. Bioscience, Biotechnology and Biochemistry, 62 (1): 66-71. Kim, H. K., Sung, M. H., Kim, H. K., and Oh, T. K., 1994. Occurrence of thermostable lipase in thermophilic Bacilus sp. Strain 398. Bioscience, Biotechnology and Biochemistry, 58 (5): 961-962. Kim, M. H., Kim, H. K., Lee, J. K., Park, S. Y., and Oh, T. K., 2000. Thermostable lipase of Bacillus stearothermophilus: high-level production, purification, and calcium-dependent thermostability. Bioscience, Biotechnology and Biochemistry, 64 (2): 280-286. Kulkarni, N., and Gadre, R. V., 1999. A novel alkaline, thermostable, protease-free lipase from Pseudomonas sp. Biotechnology Letters, 21: 897-899. Kumar, S., Kikon, K., Upadhyay, A., Kanwar, S. S., and Gupta, R., 2005. Production, purification, and characterization of lipase from thermophilic and alkaliphilic Bacillus coagulans BTS-3. Protein Expression and Purification, 41: 38-44. Lee, D. W., Kim, H. W., Lee, K. W., Kim, B. C., Choe, E. A., Lee, H. S., Kim, D. S., and Pyun, Y. R., 2001. Purification and characterization of two distinct thermostable lipases from the gram-positive thermophilic bacterium Bacillus thermoleovorans ID-1. Enzyme and Microbial Technology, 29: 363-371. Lee, D. W., Koh, Y. S., Kim, K. J., Kim, B. C., Choi, H. J., Kim, D. S., Suhartono, M. T., and Pyun, Y. R., 1999. Isolation and characterization of a thermophilic lipase from Bacillus thermoleovorans ID-1. FEMS Microbiology Letters, 179: 393-400. Leow, T. C., Rahman, R. N. Z. R. A., Basri, M., and Salleh, A. B., 2004. High level expression of thermostable lipase from Geobacillus sp. strain T1. Bioscience, Biotechnology and Biochemistry, 68 (1): 96-103. Li, H., and Zhang, X., 2005. Characterization of thermostable lipase from thermophilic Geobacillus sp. TW1. Protein Expression and Purification, 42: 153-159. Llarch, A., Logan, N. A., Castellví, J., Prieto, M. J., and Guinea, J., 1997. Isolation and characterization of thermophilic Bacillus spp. from geothermal environments on deception island, south Shetlanh Archipelago. Microbiological Ecology, 34: 58-65. Nawani, N., Dosanjh, N. S., and Kaur, J., 1998. A novel thermostable lipase from a thermophilic Bacillus sp.: characterization and esterification studies. Biotechnology Letters, 20 (10): 997-1000. Rahman, R. N. Z. R. A., Leow, T. C., Basri, M., and Salleh, A. B., 2005. Secretory Expression of Thermostable T1 Lipase through Bacteriocin Release Protein (BRP). Protein Purification and Expression, 40: 411-416. Rathi, P., Bradoo, S., Saxena, R. K., and Gupta, R., 2000. A hyper-thermostable, alkaline lipase from Pseudomonas sp. with the property of thermal activation. Biotechnology Letters, 22: 495-498. Rua, M. L., Atomi, H., Schmidt-Dannert, C., and Schmid, R. D., 1998. High-level expression of the thermoalkolophilic lipase from Bacillus thermocatenulatus in Escherichia coli. Applied Microbiology and Biotechnology, 49: 405-410. Schmidt-Dannert, C., 1999. Recombinant microbial lipases for biotechnological applications. Bioorganic and Medicinal Chemistry, 7: 2123-2130.

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Schmidt-Dannert, C., Rúa, M. L., Atomi, H., and Schmid, R. D., 1996. Thermoalkalophilic lipase of Bacillus thermocatenulatus. I. Molecular cloning, nucleotide sequence, purification and some properties. Biochimica et Biophysica Acta, 1301: 105-114. Schmidt-Dannert, C., Sztajer, H., Stöcklein, W., Menge, U., and Schmid, R. D., 1994. Screening, purification and properties of a thermophilic lipase from Bacillus thermocatenulatus. Biochimica et Biophysica Acta, 1214: 45-53. Sharma, R., Soni, S. K., Vohra, R. M., Gupta, L. K., and Gupta, J. K., 2001. Purification and characterization of a thermostable alkaline lipase from a new thermophilic Bacillus sp. RSJ-1. Process Biochemistry, 37: 1075-1084. Sigurgísladóttir, S., Konráðsdóttir, M., Jónsson, A., Kristjánsson, J. K., and Matthiasson, E., 1993. Lipase activity of thermophilic bacteria from icelandic hot springs. Biotechnology Letters, 15 (4): 361-366. Sinchaikul, S., Sookkheo, B., Phutrakul, S., and Pan, F. M., 2001. Optimization of a thermostable lipase from Bacillus stearothermophilus P1: Overexpression, purification, and characterization. Protein Expression and Purification, 22: 388-398. Sugihara, A., Tani, T., and Tominaga, Y., 1991. Purification and characterization of a novel thermostable lipase from Bacillus sp. Journal of Biochemistry, 109 (2): 211-215. Sugihara, A., Ueshima, M., Shimada, Y., Tsunasawa, S., and Tominaga, Y., 1992. Purification and characterization of a novel thermostable lipase from Pseudomonas cepacia. Journal of Biochemistry, 112: 598-603. Wang, Y., Srivastava, K. C., Shen, G. J., and Wang, H. Y., 1995. Thermostable alkaline lipase from a newly isolated thermophilic Bacillus, strain A30-1 (ATCC 53841). Journal of Fermentation and Bioengineering, 79 (5): 433-438.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 63-76

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 4

ORGANIC SOLVENT TOLERANT LIPASES Syarul Nataqain Baharum, Mohamad Ropaning Sulong, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri ABSTRACT The nature of the enzymes will be denatured in the presence of the organic solvents. Solvents not only affect enzyme stability but also change enzyme specificity. Most of the reports have been published concerning solvent stable enzymes. However, these enzymes are not naturally solvent stable enzymes that direct attempts from screening of organic solvent tolerant bacteria. In this chapter, we review the production and characterization of organic solvent tolerant lipases on the first attemps to screen organic solvent tolerant lipases from Bacillus sphaericus 205y, Bacillus sp. strain 42 and Pseudomonas sp. S5. All of these bacteria were proven to be organic solvent tolerant lipase producers. These lipases were stable in the organic solvents with log P value between 2.0 to 3.6. Organic solvent tolerant lipases from Pseudomonas sp. S5, B. sphaericus 205y, Bacillus sp. strain 42 were successfully purified to homogeneity. The optimum temperatures of these lipases were 45°C, 55°C and 70°C, respectively. These purified lipases exhibited great stability and high activity in most of the organic solvents tested, therefore, these valuable criteria should not be neglected as important products in different fields could be developed from these lipases.

INTRODUCTION The most common reason for using organic media for enzymatic reactions is because the substrate to be converted has poor solubility in water. The addition of a moderate amount of organic solvent can, in a simple way, increase the solubility of hydrophobic substrates and thereby make the reaction feasible. The presence of organic solvent may cause enzyme inactivation. The degree of activation depends on the type of solvents used and water concentration. According to Klibanov (1986),

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some solvents seem to have little effect on water molecules which are tightly bound to the enzyme, while relatively hydrophilic solvents are capable of stripping off even the essential water from the enzyme surface, leading to insufficiently hydrated enzyme molecules and subsequently decreasing the enzyme activity. The stripping of water from an enzyme by a solvent has been correlated with solvent polarity and to a lesser extent to solvent hydrophobicity (Log P). For instance, methanol desorbed about 60% of the water bound to the enzymes, while hexane only desorbed 0.5% (Klibanov, 1986). Since enzyme and solvent compete for water, the optimal water content is also related to the amount of enzyme and the concentration of substrate (Hirata et al., 1990). Removal of water from an enzyme by organic solvents can be minimized by hydrophilization of the enzyme surface. In this way, the hydration shell of the enzyme can be retained. The nature of the solvent can interact directly or indirectly with substrates and products, thus affecting the enzyme activity. Solvents can alter the concentrations of substrates or products in the aqueous layer around the enzyme, since substrates must penetrate into the water layer for the reaction to occur and products must partition out of the layer to drive the reaction forward; hence the solvent affects activity. Solvents not only affect enzymes stability but also change enzyme specificity, for instance substrate specificity, enantioselectivity, regioselectivity and chemoselectivity (Yang and Russell, 1996). Enzymes are more thermostable in non-aqueous media compared to aqueous media (Koops et al., 1999). The increase in structural rigidity and a lower rate of chemical deterioration are responsible for the high thermal stability of many proteins in dry organic solvents. Removal of water from the protein increases the strength of intramolecular hydrogen bonds and salt bridges that stabilize the proteins in their native conformation and creates a high kinetic barrier between the native and the unfolded state. Thermostability of enzymes is usually higher in hydrophobic organic solvents that are essentially inert towards the charged amino acid side chains found on protein surfaces (Zaks and Klibanov, 1984). The lyophilization of proteins from aqueous solutions at the optimal pH and the use of lyoprotectants increase the thermostability of enzymes in non-aqueous media even further.

SOURCES OF ORGANIC SOLVENT TOLERANT LIPASES Some organic solvents are highly toxic to microorganisms. However, it has been reported that Pseudomonas aeruginosa ST-001 (Aono et al., 1992), P. putida Idaho (Cruden et al., 1992), P. putida IH-2000 (Inoue and Hirokoshi, 1989), Pseudomonas sp. strain TOR (Nakajima et al., 1992) and P. aeruginosa LST-03 (Ogino et al., 1994) can grow in media containing organic solvents at a high concentration. Many different mechanisms have been described to contribute to solvent resistance, but despite these efforts no comprehensive overview is available to explain the physiological response of microorganisms to toxic organic solvents (Kieboom et al., 1998). In 1989, Inoue and Hirokoshi reported the isolation of a toluene-tolerant P. putida strain that grew in a two-phase toluene-water system. This finding came as a surprise because it had been known for a long time that toluene and other solvents such as benzene or octanol were very toxic to microorganisms.

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Kieboom et al., (1998) investigated the ability of P. putida S12 to withstand toxic concentrations of toluene and other organic solvents. This organism evolved at least two mechanisms to combat the accumulation of hydrophobic solvents in the membrane or the interior of the cell. The solvent-tolerant Pseudomonas species can be used in environmental biotechnology as well as in biotechnological production processes in two-liquid water-solvent systems (Bont, 1998).

ORGANIC SOLVENT TOLERANT LIPASES The ability of enzymes becoming active in the presence of organic solvents has received much attention in the past two decades (Sellek and Chaudhuri, 1999). Some reports have been published concerning natural enzymes which are organic solvent-stable, such as lipases in organic solvents. Ogino et al., (1994), screened for organic solvent-tolerant microorganisms which produce a lipolytic enzyme from natural sources of soil. By using olive oil as the sole source of carbon and cyclohexane as the organic solvent in an enrichment medium, organic solvent tolerant Pseudomonas aeruginosa LST-03, which secretes an organic solvent-stable lipase was isolated (Ogino et al., 1994; Ogino et al., 1999). These researchers reported that P. aeruginosa LST-03 lipase was very stable in toluene (log P= 2.5) and cyclohexane (log P= 3.1). Back then, this is the only organic solvent tolerant lipase reported, that was discovered by screening. On the other hand, some enzymes have been reported to be organic solvent stable. However, these enzymes may have not been discovered as a result of direct attempts to screen for organic solvent stable enzymes. Sharma et al., (2001) reported that their Pseudomonas sp. AG-8 was isolated from a local milk processing plant and found activated in the presence of ethanol and methanol. A lipase from P. cepacia showed higher catalytic activity in n-hexane and cyclohexane (Mine et al., 2000). In another study, a lipase from P. aeruginosa YS-7 showed good stability when incubated in 20% and 50% (w/v) of ethanol, isopropanol, ethylene glycol and propylene glycol (Shabtai and Mishne, 1992). In addition, Choo et al., (1998) reported that a lipase from Pseudomonas sp. B11-1 was activated when incubated at 25ºC for 1h in 15% (v/v) methanol, ethanol, dimetyl sulfoxide (DMSO) and n-ndimethylformamide (DMF). Lipase from Rhizopus oryzae is fairly stable in alkanes and long chain alcohols but easily denatured in hydrophobic solvents such as acetone or short chain (C1- C3) alcohol (Hiol et. al., 2000).

PRODUCTION OF ORGANIC SOLVENT TOLERANT LIPASES Many studies have been undertaken to define the optimal culture and nutritional requirements for lipase production. Lipase production is influenced by the type and concentration of carbon and nitrogen sources. Lipidic carbon sources seem to be generally essential for obtaining a high lipase yield, although a few researchers have produced good yields in the absence of fats and oils. However, there are limited studies conducted on the optimization of organic solvent tolerant lipases and almost no attempt made to study the

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optimum condition for its production. Nevertheless, many researchers have standardized the production medium with peptone and olive oil as nitrogen and carbon sources, respectively. Meanwhile, the higher initial pH was applied to the lipase production medium. The enrichment cultivation on olive oil was carried out on the assumption that microorganisms capable of growth on olive oil are capable of producing lipases; and high alkaline pH was employed on the assumption that only bacteria could grow under this condition (Gao et al., 2000). For instants, Ogino et al., (1994) reported that the medium for organic solvent tolerant LST-03 lipase production was enriched with polypeptone and yeast extract at pH 7.5 and 30ºC. The lipase production for thermostable and solvent stable extracellular lipase from Pseudomonas AG-8 was studied in medium containing glucose and yeast extract at pH 7.5 (Sharma et al., 2001). Meanwhile, Gao et al., (2000) reported that the optimum production of nonaqueous enzymatic catalysis of lipase from a newly isolated Pseudomonas strain was at pH 9.0, 45ºC with olive oil as a substrate. The production medium for another Pseudomonas lipase was composed from ground soybean, corn steep liquor and soluble starch and was adjusted to pH 9.0 and the lipase producer was incubated at 30ºC for 72 h (Dong et al., 1999).

PURIFICATION AND CHARACTERIZATION OF ORGANIC SOLVENT TOLERANT LIPASES The objective of purification of an enzyme is to get rid of as much unwanted proteins as possible while retaining the enzyme activity. Purification of the protein leads to better understanding on the effect of organic solvent on the enzyme activity and stability. Dong et al., (1999) suggested the employment of crude enzyme preparation to elucidate the properties of lipase in nonaqueous media, but appeared unlikely to obtain positive results owing to the unpredictable effects of the impurities in the lipase. An organic solvent stable lipase from Pseudomonas aeruginosa LST-03 was purified to 34.7 fold with an overall yield of 12.6% by ammonium sulphate precipitation, ion exchange and hydrophobic interaction chromatography (Ogino et al., 2000). This organic solvent tolerant lipase exhibited molecular mass of 27.1 kDa by SDS-PAGE and 36 kDa by gel filtration. Meanwhile, other thermostable and solvent stable extracellular lipase from Pseudomonas AG-8 was partially purified to 2.1 fold and 90% of recovery using 60% ammonium sulphate precipitation. But the molecular mass of the lipase was not reported (Sharma et al., 2001). An extracellular lipase from other Pseudomonas sp. was purified to homogeneity by extraction, Bio-gel P-10 chromatography and Superose 12B chromatography with overall yield of 64% and 37 purification fold. The size of the purified enzyme was 30 kDa (Dong et al., 1999). Meanwhile, Shabtai and Mishne, (1992) reported that the P. aeruginosa YS-7 lipase was purified to homogeneity using acetone precipitation and hydrophobic interaction chromatography with 49% of recovery and 1333 U/mg of specific activity.

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STABILITY IN ORGANIC SOLVENT Enzymes in organic solvents have been largely studied and employed in the areas of product synthesis, food production and even biochemical analysis. The use of organic solvents is especially advantageous to transform substrates that are unstable or poorly soluble in water (Secundo and Carrea, 2002). However, enzymes are generally not stable in the presence of organic solvents and are apt to denature, which has led to the development of several methods for stabilizing them. Nevertheless, enzymes that remain stable in the presence of organic solvents without the need for special stabilization measures could be very useful in industry (Ogino et al., 2000). Therefore, many researchers have investigated the stability of lipase in organic solvents. Organic solvent stable purified lipase from organic solvent tolerant Pseudomonas aeruginosa LST-03 was stable in the presence of n-decane, DMSO, n-octane, n-heptane, isooctane and cylohexane (Ogino et al., 2000). In 1990, Iizumi et al., reported that the purified lipase from Pseudomonas species KWI-56 was stable against organic solvents such as acetone, methanol and ethanol. P. cepacia lipase was inactivated in presence of n-propanol and pyridine (Sugihara et al., 1992). Jonzo et al., (2000) reported that an increased ester yield by Candida rugosa lipase was observed at about 60% with hexanol and octanol. The stability of the Aspergillus terreus lipase was affected by ethanol at 90% concentration, and propan-2-ol above 20%, had drastic inhibitory effects on lipase stability (Yadav et al., 1998). A purified lipase from Mucor hiemalis f. hiemalis showed a good stability in water-immiscible organic solvents like hexane, heptane and isooctane. Acetonitrile at a concentration of 5% (v/v) slightly activated the enzyme activity (Hiol et al., 1999). In 2000, Hiol et al., reported that the enzyme from Rhizopus oryzae was fairly stable in alkanes and long chain alcohols but it is easily denatured in hydrophilic solvents such as acetone or short chain (C1-C2) alcohol.

EFFECT OF pH ON LIPASE ACTIVITY AND STABILITY Many researchers reported on Pseudomonas lipases that were more active in alkaline pH. An organic solvent tolerant Pseudomonas lipase LST-03 has maximum activity at pH ranging from pH 6.0 to 10.0 and showed great stability at pH range 5.0-8.0 for only 10 min at 30°C (Ogino et al., 2000). Meanwhile, AG-8 lipase still retained more than 90-100% activity in pH 7.0 to 10.0 for 24 hours at 25°C (Sharma et al., 2001). Lipases from Pseudomonas cepacia and Pseudomonas sp. KWI-56 showed an optimum pH at 5.5 to 6.5 and pH 5.5 to pH 7.0, respectively (Iizumi et al., 1990; Sugihara et al., 1992). However, most of the lipases from fungal origin showed stability on broad range of pH. For instance, Aspergillus terreus organic solvent stable lipase has shown good pH tolerance (3.0-12.0) and was stable over a pH range of 4.0-10.0 for 24 hours (Yadav et al., 1998). Hiol et al., (1999) reported that the optimum pH for Mucor hiemalis f. hiemalis lipase was 7.0 and stable in the pH range of 4-9.

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EFFECT OF TEMPERATURE ON LIPASE ACTIVITY AND STABILITY Many researchers have determined the optimum temperature for their organic solvent stable lipases. Most of the lipases reported were stable at higher temperature. The optimum temperature for purified lipase from Pseudomonas sp. KWI-56 was 60°C and highly thermostable, retaining 96% activity at 60°C for 24 hours, 76% at 70°C after 3 hours (Iizumi et al., 1990). Sharma et al., (2001) reported that their Pseudomonas sp. AG-8 lipase had optimal activity at 45°C. This lipase had a half life of 50 min at 67°C. Meanwhile, Sugihara et al., (1992) had purified Pseudomonas cepacia lipase that showed optimal temperature at 5560°C. In 1991, Sugihara et al., reported that the optimal temperature for purified lipase from Bacillus sp. was 60°C and stable up to 65°C for 30 min at pH 5.6. Meanwhile, Bacillus thermocatenulatus lipase was stable up to 40ºC for 30 min (Dannert et al., 1994). In contrast, an organic solvent stable LST-03 lipase showed higher stability at a lower temperature. This organic solvent lipase had maximum activity at 37°C (Ogino et al., 2000) and showed a stability below 40°C, for 10 min.

SUBSTRATE SPECIFICITY According to Ogino et al., (2000) lipases had highest activity against coconut oil because the principal fatty acid component was lauric acid (12:0) which was more hydrolyzable by the lipase than oleic acid (18:1), the principal fatty acid component of olive oil. In the oleochemical industry, enzymatic hydrolysis of oils and fats is an energy-saving process, compared to conventional high-temperature and high-pressure processes (Iizumi et al., 1990). From the viewpoint of degree of hydrolysis, lipases which hydrolyze longer carbon chain lengths of substrates are better candidates for this purpose. Similarly, the purified lipases by Pseudomonas sp. hydrolyzed various oils and fats efficiently such as palm oil, soybean oil and coconut oil (Iizumi et al., 1990; Jinwal et al., 2003). Meanwhile, Ogino et al., (2000) reported that LST 03 lipase had highest hydrolytic activity against tricaproin (C6).

EFFECT OF METAL IONS ON ENZYME STABILITY According to Voet et al., (1999), nearly one-third of all enzymes require the presence of metal ions for catalytic activity. This group of enzymes includes the metalloenzymes, which contain tightly bound metal ion cofactors, most commonly transition metal ions such as Fe2+, Fe3+, Zn2+, Mn2+, Cu2+ or Co2+ . Metal-activated enzymes, in contrast, loosely bind metal ions from solution, usually the alkali and alkaline earth metal ions such as Na+, K+, Mg2+or Ca2+. In this group of enzymes, the ions often play a structural rather than a catalytic role. Lipases show variable stability towards various metal ions, but most preferred Calcium ions as stabilizer. Sharma et al., (2001), reported that Pseudomonas sp. lipase AG-8 was activated in the presence of Ca2+, while Fe3+ and Zn2+ strongly inhibited the activity. The activity of P.

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cepacia was inhibited by Cu2+ and Hg2+, whereas Ca2+, Ni2+, Sr2+ and Co2+ and EDTA did not affect the enzyme activity (Sugihara et al., 1992). Sugihara et al., (1991) tested the effect of metal ions to the purified lipase from Bacillus sp. Cu2+, Zn2+ and Hg2+ showed 70% inhibition whereas Mg2+, Ca2+, Fe2+, Zn2+, Sr2+, Sn2+ and Ba2+ and EDTA did not significantly affect the enzyme activity. Hiol et al., (1999) reported that Mg2+, Ca2+, Co2+, Mn2+ and Na+ enhanced lipase activity from Mucor hiemalis f. hiemalis, whereas Fe2+, Ba2+ and Cu2+ strongly reduced the lipase activity. The enzyme activity was not affected by EDTA, PMSF and Benzamidine. The activity of Aspergillus terreus lipase was stimulated by Mg2+ and Ca2+ ions, whereas Co2+, Cu2+ Fe2+ and Ni+ ions cause its inhibition (Yadav et al., 1998).

OUR WORK In the past, most microorganisms were considered as the causative agents for various diseases. Today, the societies’ view towards microorganisms has significantly changed. The progress in biotechnology particularly in the field of industrial microbiology has led to many researchers to take the economic opportunities in the production of numerous valuable products. Lipases are among such valuable products. Therefore, scientists began to isolate and screen various microorganisms, mainly bacteria, in the search to discover better lipases. Based on the multiple criteria, lipases have been categorized into different types such as thermostable lipase, alkaline lipase and organic solvent tolerant lipase. With respect to industrial applications, organic solvent tolerant lipases are the more promising types due to the many advantages that could be derived. In this chapter, we review the screening and isolation, growth and production, purification and characterization of organic solvent tolerant lipases produced by different types of bacteria such as Bacillus sphaericus 205y, Bacillus sp. strain 42 and Pseudomonas sp. S5.

Isolation and Screening of Organic Solvent Tolerant Lipase Producer Different genus of organic solvent tolerant lipase bacteria, known to be BTEX (Benzene, Toluene, Ethyl-Benzene and Xylene) degraders, have been isolated and screened for lipolytic activity. Pseudomonas sp. S5 was locally isolated from soil sample at Serdang workshop area (Baharum et al., 2003), Bacillus sphaericus 205y was isolated from polluted area in Port Dickson (Hun et al., 2003) and Bacillus sp. strain 42 was isolated from palm oil mill effluents in Johor (El Taweel et al., 2005). All of the bacteria were proven to be organic solvent tolerant lipase producers. Lipolytic screening of the lipases involved two assays; the plate and the broth assays. All the organic solvent tolerant lipase producers were screened for lipase activity on tributyrin and triolein agar plates (Samad et al., 1998), where a halo on tributyrin and intensive blue colour on triolein indicates its detection. While for the broth assay, the lipase activity was assayed according to the method by Kwon and Rhee (1986). Both the plate and broth assays have indicated positive lipase production.

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Effect of Physical Factors and Nutritional Factors on Lipase Production The growth and lipase production will depend mainly on the physical as well as nutritional factors; therefore, different media are usually required for growth and production of organic solvent tolerant lipase producers. The strain S5 of Pseudomonas sp. has a very simple nutritional requirement with neutral pH and at a temperature within the mesophilic range. Bacillus sphaericus 205y grows better on liquid media containing nutrient broth, gum arabic, CaCl2.2H2O, Tween 80 and Olive oil while for Bacillus sp. strain 42, the optimum medium for growth contains olive oil, tryptone, yeast extract and some metal ions. The optimum lipase production is at static condition at 37˚C for Pseudomonas sp. S5 hence not requiring any shaking, while a shaking rate of 150 rpm at 37˚C and 50°C were the best for lipase production for B. sphaericus 205y and Bacillus sp. strain 42, respectively. On the other hand, 6% (v/v) of inoculum size was found to be the best for the growth of Pseudomonas sp. S5, subsequently the highest lipase production as well as activity was obtained.

Effect of Various Organic Solvents on Crude Lipase Stability Crude enzymes from isolates S5 and 205y were tested on their organic solvent stability. Fourteen organic solvents with different log P were used. According to Wu et al., (1997) the polarity of organic solvents can be measured with value of log P between water and 1octanol. Table 1 listed the log P value (Laane et al., 1986) of various organic solvents used in this study. Isolates 205w and S5 were incubated in various organic solvents (25%) for 30 min at 150rpm and 37ºC before remaining lipase activities determined. These results showed that the S5 lipase was stable in most of the organic solvents with log P value of 2.0 to 3.6. Benzene and 1-octanol with log P of 2.0 and 2.9 respectively enhanced the lipase stability as compared with the culture in the absence of organic solvents (Baharum et al., 2003). Meanwhile lipase from isolate 205y was stable in p-xylene and nhexane. The enzyme suffered detrimental effect in low log P solvents such as dimethyl sulfoxide and acetonitrile. Approximately 80% activity was lost in the presence of dimethyl sulfoxide and complete deactivation in the presence of acetonitrile (Hun et al., 2003). The higher lipase stability was exhibited by both isolates in the presence of n-hexane. No lipase activity was detected in n-hexadecane, which has high log P value of 8.8. Basri et al., (1997) also found that solvent of high log P values have detrimental effect on lipase activity. This may due to the relatively high viscosity of the solvents, which hindered efficient interaction between enzymes and substrates.

Purification of Organic Solvent Tolerant Lipases For any crude biocatalysts or proteins to be used for further downstream applications, purification is becoming a prerequisite to ensure the purity as well as the safety of these proteins. The degree of the purity usually depends on the application of the end products; for example, the highest degree of the enzyme purity (not less 99%) have to be use in pharmaceutical application. Recently, various purification methods have been recognized; as no one particular method can be applied for the purification of the proteins since different

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proteins may require different purification techniques. Hence, all the organic solvent tolerant lipases producers: Pseudomonas sp. S5, Bacillus sphaericus 205y and Bacillus sp. strain 42 have been successfully purified by different methods. Table 1: Effect of Various Organic Solvents on Crude Lipase Stability Organic Solvents at concentration of 25%

No solvent Dimethyl sulfoxide Acetonitrile Ethyl acetate 1-pentanol Benzene 1-heptanol Toluene 1-octanol p-xylene Cyclohexane n-hexane 1-decanol n-decane n-dodecane n-tetradecane n-hexadecane

Log P

-1.22 -0.15 0.68 1.3 2.0 2.4 2.5 2.9 3.1 3.2 3.6 4.0 5.6 6.6 7.6 8.8

Relative activity (%)

S5* 100 n/a n/a 25 98 170 45 110 160 100 275 420 0 0 30 15 15

205y† 100 19 0 n/a n/a n/a n/a n/a n/a 288 n/a 349 n/a n/a n/a n/a 0

S42‡ 100 n/a n/a 46.4 n/a 105 n/a 84.7 n/a 79.7 n/a 104 90 n/a 70.9 94.5 104

Three milliliters of cell-free supernatant of the culture was incubated with one milliliters of organic solvents at 37ºC, 100rpm shaking for 30 min. The remaining lipase activity relative to the nonsolvent containing control was measured.

An organic solvent tolerant lipase from Pseudomonas sp. S5 was purified to homogeneity using Con A-Sepharose column chromatography (Amersham-Pharmacia Biotech, Uppsala, Sweden) followed by ion-exchange chromatography on DEAE-Sephacel (Rahman et al., 2005). The lipase was purified to 387-fold purification and was showed to have a molecular mass of 60 kDa on SDS-PAGE (Figure 1) while another organic solvent tolerant lipase from B. sphaericus 205y was purified using ultrafiltration followed by hydrophobic interaction chromatography (HIC) (Amersham-Pharmacia Biotech, Uppsala, Sweden) and the organic solvent tolerant Bacillus sp. strain 42 was purified as one step purification using affinity column chromatography on Ni-NTA.

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kDa 116 97.2 66.4 Purified S5 lipase 55.6 48.0

36.5

26.6

14.3

1

2

3

4

Figure 1: Electrophoresis of strain S5 Lipase in 10% Polyacrylamide Gel Under Denaturing Conditions. Lane 1: Marker, Lane 2: Crude enzyme, Lane 3: Affinity chromatography, Lane 4: Ion Exchange (purified S5 lipase) (Source: Rahman et al., 2005)

Characterization of Organic Solvent Tolerant Lipase The purified organic solvent tolerant lipase from Pseudomonas sp. S5 showed an optimum lipase activity at pH 9.0 and was stable at pH range 6.0 to 9.0. In contrast, the optimum pH for lipase activity of the purified organic solvent tolerant lipase from B. sphaericus 205y as well as Bacillus sp. strain 42 was found at neutral pH and have a wide range of pH stability. The effect of temperature as well as the thermostability test on the lipases exhibited that Pseudomonas sp. S5, B. sphaericus 205y and Bacillus sp. strain 42 lipase have the optimum lipase activity at 45°C, 55°C and 70°C respectively. The half-life for both B. sphaericus 205y

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and Bacillus sp. strain 42 were more than 30 min at temperature above 45°C therefore are considered as a thermostable lipase. The study on the effect of reducing agents exhibits interesting results; both oxidizing and reducing agents have decreased the activity of these lipases (205y, S5 and 42). The metal chelator agent, 1mM of EDTA, has no effect on all the three lipases, indicating that these lipases are not metalloenzyme type. However, PMSF has an effect on the lipases, where 1 mM of PMSF decreased the activity of all lipases, indicating the lipases are of the serine hydrolase type (unpublished data). Metal ions such as Ca2+ and Mg2+ have stimulated the lipases activities. The study on substrate specificity showed that palm oil as a natural oil and triolein as a synthetic triglyceride have stimulated the lipase activity of Pseudomonas sp. S5 while both B. sphaericus 205y and Bacillus sp. strain 42 lipases hydrolysed medium chain length triglycerides. Finally, all the three purified lipases from Pseudomonas sp. S5, B. sphaericus 205y and Bacillus sp. strain 42 exhibited great stability and high activity in most of the organic solvents tested, therefore, these valuable criteria should not be neglected as important products in different fields could be developed from these lipases. Table 2 summarizes the characteristic of pure lipases from isolates S5, 205y and 42. Table 2: Characteristics of organic solvent tolerant lipases Characteristic of pure lipases Temperature

pH

Substrate Metal ion Stability in Organic Solvent

S5

205y

42

45°C, half life 1h at 50°C

55°C, 30 min at temperature above 45°C pH optimum: pH 7.0

70°C, 30 min at temperature above 45°C pH optimum: pH 7.0

C8- C12 Ca and Mg2+ Stable in ethanol, 1decanol and nhexadecane. Enhanced in DMSO, methanol, p-xylene and n-decane

C8- C12 Ca and Mg2+ Stable in propylacetate, heptanol and octanol

pH optimum: pH 9.0, stable at pH range 6.0 to 9.0 C12- C18 Ca2+ and Mg2+ Stable in 1-pentanol, chloroform, n-dodecane, 1-octanol and cyclohexane for 2h

2+

2+

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Jonzo, M. D., Hiol, A., Zagol, I., Druet, D., and Comeau, L. C., 2000. Concentrates of DHA from fish oil by selective esterification of cholesterol by immobilized isoforms of lipase from Candida rugosa. Enzyme and Microbial Technology, 27: 443-450. Kieboom, J., Jonathan, J. D., De Bont, J. A. M., and Zylstra, G. J., 1998. Identification and molecular characterization of an efflux pump involved in Pseudomonas putida S12 solvent tolerance. Journal of Biological Chemistry, 273: 85-91. Klibanov, A. M., 1986. Enzymes that work in organic solvents. Chemical Technology, 16: 354-359. Koops, B. C., Verheij, H. M., Slotboom, A. J., and Egmond, M. R., 1999. Effect of chemical modification on the activity of lipases in organic solvents. Enzyme and Microbial Technology, 25: 622-631. Kwon, D. Y., and Rhee, J. S., 1986. A simple and rapid colorimetric method for determination of free fatty acids for lipase assays. Journal of American Oil Chemists’ Society, 63: 29-32. Laane, C., Bocren, S., Vos, K., and Veeger, C., 1987. Rules of optimization of biocatalysis in organic solvent. Biotechnology and Bioengineering, 30: 81-87. Mine, Y., Fukunaga, K., Maruoka, N., Nakao, K., and Sugimura, Y., 2000. Preparation of detergent-lipase complexes utilizing water-soluble amphiphiles in single aqueous phase and catalysis of transesterifications in homogeneous organic solvents. Journal of Bioscience and Bioengineering, 90 (6): 631-636. Nakajima, H., Kobayashi, H., Aono, R., and Horikoshi, K., 1992. Isolation and identification of toluene-tolerant Pseudomonas strains. Bioscience, Biotechnology and Biochemistry, 56 (11): 1872-1873. Ogino, H., Miyamoto, K., and Ishikawa, H., 1994. Organic solvent- tolerant bacterium which secretes organic solvent-stable lipolytic enzyme. Applied and Environmental Microbiology, 60 (10): 3884-3886. Ogino, H., Miyamoto, K., Yasuda, M., Ishimi, K., and Ishikawa, H., 1999. Growth of organic solvent-tolerant Pseudomonas aeruginosa LST-03 in the presence of various organic solvents and production of lipolytic enzyme in the presence of cyclohexane. Biochemical Engineering Journal, 4: 1-6. Ogino, H., Nakagawa, S., Shinya, K., Muto, T., Fujimura, N., Yasuda, M., and Ishikawa, H., 2000. Purification and characterization of organic solvent-stable lipase from organic solvent-tolerant Pseudomonas aeruginosa LST-03. Journal of Bioscience and Bioengineering, 89 (5): 451-457. Rahman, R. N. Z. A., Baharum, S. N., Basri, M., and Salleh, A. B., 2005. High-yield purification of an organic solvent-tolerant lipase from Pseudomonas sp. S5. Analytical Biochemistry, 341: 267-274. Samad, M. Y. A., Razak, C. N. A., Salleh, A. B., Yunus, W. M. Z. W., Ampon K., and Basri, M., 1998. A plate assay for primary screening of lipase activity. Journal of Microbiological Methods, 9: 51-56. Secundo, F., and Carrea, G., 2002. Lipase activity and conformation in neat organic solvents. Journal of Molecular Catalysis B: Enzymatic, 19-20: 93-102. Sellek, G. A., and Chaudhuri, J. B., 1999. Biocatalysis in organic media using enzymes from extramophiles. Enzyme and Microbial Technology 25: 471-482.

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Shabtai, Y., and Mishne, N. D., 1992. Production, purification and properties of a lipase from a bacterium (Pseudomonas aeruginosa YS-7) capable of growing in water-restricted environments. Applied and Environmental Microbiology, 58(3): 174-180. Sharma, A. K., Tiwari, R. P., and Hoondal, G. S., 2001. Properties of a thermostable and solvent stable extracellular lipase from a Pseudomonas sp. AG-8. Journal of Basic Microbiology, 41 (6): 363-366. Sugihara, A, Tani, T., and Tominaga, Y., 1991. Purification and characterization of novel thermostable lipase from Bacillus sp. Journal of Biochemistry, 109: 211-219. Sugihara, A., Ueshima, M., Shimada, Y., Tsunasawa, S., and Tominaga, S., 1992. Purification and characterization of novel thermostable lipase from Pseudomonas cepacia. Journal of Biochemistry, 112: 598-608. Voet, D., Voet J. G., and Pratt C. W., 1999. Fundamentals of Biochemistry. John Wiley and Sons. Inc. New York. pp: 295. Wu, W. H., Akoh C. C., and Philips, R. S. 1997. Stereoselective acylation of DL- methol in organic solvents by an immobilized lipase from Pseudomonas cepacia with vinyl propionate. Journal of American Oil Chemists’ Society, 74 (4): 435-439. Yadav, R. P, Saxena, R. K., Gupta, R., and Davidson, W. S., 1998. Purification and characterization of a regiospecific lipase from Aspergillus terreus. Biotechnology and Applied Biochemistry, 28: 243-249. Yang, Z., and Russell, A. J., 1996. Fundamentals of Non-aqueous Enzymology, Enzymatic Reaction in Organic Media. Ed. Koskinen, A.M.P., and A. M. Klibanov. Blackie Academic and Professional. London. pp: 55-57. Zaks, A., and Klibanov, A. M., 1984. Enzymatic catalysis in organic media at 100ºC. Science, 224: 1249-1251.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 77-93

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 5

THERMOSTABLE PROTEASES Noor Azlina Ibrahim, Thean Chor Leow, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri ABSTRACT New proteases are constantly isolated. Proteases, mainly the alkaline proteases are widely used in the detergent formulation. Thermostable alkaline proteases are of great interest in the detergent industry because these enzymes are stable and able to retain its activities over broad range of pH and temperatures. Most thermostable alkaline proteases reported are from Bacillus spp. In this chapter, we introduce a new thermostable alkaline protease produced from Bacillus stearothermophilus strain F1. The protease production requirements by strain F1, the properties and characterization of this enzyme are discussed. The protease F1 gene was successfully cloned and expressed into E. coli XL1Blue. The bacteriocin release protein (BRP) system was utilized; to release the recombinant F1 protease into the culture medium. The purified native and recombinant F1 protease showed a pH optimum of 9.0 and thermostabilities ranging from a half-life of 25 min at 90oC to a half-life of 4 h at 85oC. The optimum temperatures were 85oC and 80oC respectively.

INTRODUCTION Microbial proteases dominate the commercial enzyme applications, where a large market share is taken by subtilisin proteases from Bacillus spp. mainly as additive in the production of laundry detergent. An important enzyme characteristic required for such commercial applications is its thermal stability, because thermal denaturation is the common cause of enzyme inactivation. There have been a number of recent efforts to improve the thermostability of the enzymes within the currently limited knowledge on protein engineering (Suzuki et al. 1989).

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SOURCES OF THERMOSTABLE PROTEASES Organisms belonging to the genus Bacillus produce most of the commercial proteases, mainly the neutral and alkaline proteases (Rao et al., 1998). Bacillus spp. are prolific producers of extracellular hydrolytic enzymes, and in particular for the production of subtilisin proteases which has been extensively exploited for use in laundry detergents and for other applications (Godfrey and West, 1996). The strains used to produce enzymes for laundry detergents have been selected for the ability of their proteases to withstand the harsh conditions, prevailing during the wash cycles and for the high yield of this enzyme produced in industrial fermentors (Peek et al., 1993). Some thermophilic Bacillus spp. that produced thermostable proteases have been isolated. The most notable thermostable neutral metalloprotease produced was thermolysin from B. thermoproteolyticus. The optimum temperature was found to be 80oC with a half-life of 1 h at 80oC (Ohta et al., 1966). The neutral proteases from Bacillus stearothermophilus strain NCIB 8924 (Sidler and Zuber, 1972) and strain NRRL B-3880 (Sidler and Zuber, 1977) were stable up to 65oC and 70oC respectively. B. stearothermophilus strain MK 232 was reported to produce protease with optimum activity at 70oC (Kubo et al., 1988). Bacillus sp. strain AH-101 was reported to produce alkaline serine protease with optimum temperature of 80oC, in the presence of 5mM Ca2+. The optimum temperature and thermostability of most alkaline serine proteases were stabilized significantly by Ca2+. Protease HS which was stable at 60oC (Durham et al., 1987) and protease BYA were exceptional because these alkaline serine protease enzymes were reported to be thermostable in the absence of Ca2+. An alkaline protease produced by Bacillus sp. B18 was reported to have an optimum temperature and pH at 85oC and pH 12-13 respectively (Fujiwara et al, 1993). The most recent heat stable protease producer reported was B. substilis PE-11; the enzyme was most active at 60oC and pH 10, with casein as substrate (Adinarayana et al., 2003). The enzyme was almost 100% stable at 60oC for 350 min of incubation. The first reported on isolation of heat stable protease from Pseudomonas sp. was by Stepaniak et al. (1982). The protease isolated from Pseudomonas fluorescens AFT36 was reported to be stable at 80oC. Protease from P. fluorescens T16 was also found to be thermostable, retaining about 8% activity after being treated at 120oC for 10 min (Patel et al., 1983). Thermopsin, an acid protease from Sulfolobus acidocaldarius (Fusek et al., 1990; Lin and Tang, 1990) demonstrated an optimal activity at 76oC. Two strains of actinomycete, Thermomonospora fusca, have been reported to produce thermostable protease. T. fusca strain A29 and strain YX were producers of protease with optimum temperatures of 65oC and 80oC respectively (Desai and Dhala, 1969, Gusek and Kinsella, 1987). Several serine type proteases have been isolated from extreme thermophilic Thermus isolates. Caldolase from Thermus sp. strain Tok 3 (Saravani et al., 1989), aqualysin 1 from T. aquaticus (Matsuzawa et al., 1988) and Rt41A protease (Peek et al., 1992) were chelator resistant, whilst caldolysin from strain T 351 (Cowan and Daniel, 1982) and aqualysin 11 from T. aquaticus (Matsuzawa et al., 1983) were chelator positive. Aqualysin 1 was a serine protease exhibiting maximum activity between 70oC and 80oC (Matsuzawa et al., 1988). Extremely thermostable serine proteases are produced by the hyperthermophilic archaea like Thermococcus stetteri and Pyrococcus furious (Voorhorst et al., 1997). Both archaea

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produced serine proteases, which are optimally active at 85°C and 100°C, respectively. Other archaea species such as T. celer, T. litoralis and the hyperthermopilic eubacterium Thermobacteroides proteolyticus are also producers of heat stable serine type proteases (Klingeberg et al., 1991). The optimum temperatures for the first two strains were 95oC and 85oC respectively. Table 1 shows a few more thermostable proteases producers. Table1: Sources and properties of thermostable proteases Type

Source

Half life

Reference

Serine protease Pyrolysin Stetterlysin Tk subtilisin

Aquifex pyrophilus Pyrococcus furiosus Thermococcus stetteri Thermococcus kodakararaensis KOD1 Bacillus sp. Thermoactinomyces vulgaris Bacillus sp.

6 h/105ºC 4 h/100ºC 2.5 h/100ºC 20 min/90ºC

Choi et al. (1999) Voorhorst et al. (1997) Voorhorst et al. (1997) Kannan et al. (2001)

Ak.1 protease Thermitase WF146 protease

1.7 h/85ºC 1 h/85ºC 30 min/80ºC

MacIver et al. (1994) Meloun et al. (1985) Wu et al. (2004)

GROWTH AND PRODUCTION OF THERMOSTABLE PROTEASES Production of proteases from plants and animals is a time-consuming process and it is affected by political and agricultural policies. Microorganisms represent an excellent source of enzymes owing to their broad biochemical diversity and their susceptibility to genetic manipulation. Microbial proteases account for approximately 40% to 65% of the total worldwide enzyme sales (Godfrey and West, 1996, Rao et al., 1998). Nesterenkonia sp. AL-20 and Bacillus pseudofirmus AL-89 showed good growth and protease production in alkaline medium containing chicken feather as carbon and nitrogen sources. Complete solubilisation of feather was observed after 48-60 h (Gessesse et al., 2003). Maximum growth and protease production by B. subtilis PE-11 was observed at 48h in medium containing glucose, peptone and salt solution (Adinarayana et al., 2003). B. brevis utilized several carbon sources for the production of protease. Lactose was the best substrate, followed by glucose and sucrose. Among the various organic nitrogen sources, soybean meal was found to be the best. The protease was stable at 25°C for 288 h whereas, at 50 and 60°C, the half lives were 60 and 7 h respectively (Banarjee et al., 1999). Study by Fujiwara and Yamamoto (1987) showed that glucose and starch were the best carbon sources for protease production by Bacillus sp. No 221, while lactose and sucrose were least effective. Fujiwara et al., (1993) also reported glucose (1%, w/v) as the best carbon source for protease production by Bacillus sp. B18.

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PURIFICATION OF NATIVE AND RECOMBINANT THERMOSTABLE PROTEASES Numerous methods have been reported for the purification of microbial proteases. The initial step of enzyme purification mainly involves the salting out of the proteins, using organic solvents or salts at high concentrations. Ultrafiltration was commonly used to concentrate diluted protein solution prior to chromatographic analysis in protease purification (Manachini et al., 1988). Liquid chromatography methods, such as ion exchange, gel filtration, hydrophobic and affinity chromatographies were used for the subsequent steps in protease purification. A combination of gel filtration with an ion exchange chromatography was reported to purify some microbial proteases (Tsuboi et al., 1987). Van Noort et al. (1991) demonstrated that the Bacillus antibiotic, bacitracin could be used as affinity ligand to purify a protease from Aspergillus niger. Bacitracin has been shown to bind all four major mechanistic classes of proteases. MacIver et al. (1994) has successfully bound the Ak1 serine protease to bacitracin column and provided a simple and effective method for protease purification to near homogeneity. Heat-treatment was also used to purify a thermostable aminopeptidase followed by hydrophobic chromatography (Schalk et al. 1992).

CHARACTERIZATION OF PURIFIED THERMOSTABLE PROTEASES Bacterial alkaline proteases are characterized by their high activity at alkaline pH, e.g., pH 10, and their broad substrate specificity. Their optimal temperature is around 60°C (Rao et al., 1998). An extracellular subtilisin like-protease from Bacillus sp. WF146 was successfully cloned and expressed into Escherichia coli strain BL21 (DE3) (Wu et al., 2004). The optimum temperature of the purified recombinant WF146 protease was 85°C, with a half-life of 30 min at 80°C in the presence of 10mM CaCl2. Protease produced by Nesterenkonia sp. was found to be optimally active at 70°C and in the pH range of 7.0-11.0. Furthermore, unlike other alkaline proteases, the enzyme activity was found to be independent of the presence of calcium ions (Gessesse et al., 2003). The alkaline protease isolated from Bacillus subtilis PE-11 was found to be a thermostable serine protease. The purified enzyme was most active at 60°C, pH 10, with casein as a substrate. It was stable between pH 8 and 10. This enzyme was almost 100% stable at 60°C even after 350 min of incubation (Adinarayana et al., 2003).

OUR WORK Screening of Thermostable Proteases The potential use of thermostable enzymes in a range of biotechnological applications is widely acknowledged. Their inherent stability at elevated temperatures derived through directed evolution at such high growth temperatures. From a comprehensive screenings

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programmed for thermophilic proteolytic bacteria, 12 out of 100 isolates obtained from the various sampling sites were able to grow at 70°C and gave positive results on skim milk agar (SMA), indicating that they were thermophilic proteolytic bacteria (Razak et al., 1995). The casein in the skim milk can be hydrolyzed by protease. The formation of the clearing zone around the colonies indicated that the bacterium has the capacity to produce protease (Figure 1). These isolates were also grown in liquid media at different pH for quantitative screening. Isolate F1 was selected for further study based on its high protease activity and its capability to produce protease under alkaline pH medium. Furthermore, preliminary studies on the crude protease produced by isolate F1 showed that the crude enzyme was thermoresistant, retaining 100% of its activity at 70°C for 24 h. Isolate F1, that produced an extremely thermostable alkaline protease was isolated from decomposed oil palm branches (Figure 2). Based on the physical and biochemical characteristics and confirmed by API CHB identification kit, the bacterium was identified as Bacillus stearothermophilus strain F1 (Razak et al., 1995).

Figure 1. Screening of proteolytic microorganism. Isolates were isolated on skim milk agar. The ability of the bacteria to produce protease was determined by growing it on skim milk agar plate overnight at 60°C.

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Figure 2. Bacillus stearothermophilus F1 was isolated from decomposed palm oil branches.

Growth and Production of Thermostable Protease Growth parameters, either physically or nutritionally, significantly affect the growth of microorganisms. Therefore, considerable changes in fermentation media have to be made to increase the protease yield. The growth and production rate varies among organisms since the enhancing effect for one organism may be an inhibitory effect to others. Generally, extracellular protease biosynthesis requires both catabolic depression and substrate induction. An appropriate combination of physical and nutritional factors may provide profound effect in the growth of microorganism , hence the production of the enzyme.

Wild-type Protease Effect of Physical Factors on Protease Production Bacillus stearothermophilus F1 was able to secrete extracellular protease over a broad pH range (pH 5 to 12). However, the highest protease production was detected when B. stearothermophilus F1 was cultivated in an alkaline medium (pH 10). Meanwhile, the temperature range for growth extended from 50°C to 80°C in shaking cultures with 70°C being the optimum growth temperature. Protease production however was detected only up to 65°C. Protease production for strain F1 was detected after 12 h incubation with a maximum production after 24 h when grown in liquid medium (pH 10.0) at 60°C (Figure 3). This corresponded to the mid-exponential growth phase of bacteria.

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108

107

106

105

104

103

Figure 3: Growth curve of Bacillus stearothermophilus F1 in peptone medium (pH 10.0) grown at 60°C and 100 rpm agitation, showing bacteria count ( ), protease activity ( ), pH ( ) and OD679nm ( ). S denotes first observation of spores within vegetative cells. (Source: Rahman et al., 2003)

The onset of protease production coincided with the onset of sporulation, suggesting a possible correlation between sporulation and protease production in these bacteria. Shaking the culture at various shaking rates stimulated the protease production. Maximum production was achieved at 100 rpm. Higher shaking rates reduced the protease yield. Agitation had stimulated the F1 protease production. It could be due to the transfer rates of oxygen, nutrient and product from the medium to cell were enhanced and vice versa (Razak et al., 1995).

Effect of Nutritional Factors on Protease Production Most proteolytic bacteria showed a variation in their requirement for both nitrogen and carbon sources for growth as well as enzyme production. The growth and production rates could be correlated to each other, but good growth might not necessarily meant high enzyme productions. Therefore, the growth of Bacillus stearothermophilus F1 as well as its ability to produce protease in liquid media was examined in the presence of various nitrogen and carbon sources (Rahman et al., 2003). Almost all of the organic nitrogen sources tested supported luxuriant bacterial growth, but protease activity was only detected in a medium

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containing peptone, soytone, corn steep liquor, casein, gelatine and beef extract. Maximum protease production was obtained when peptone from soybean was used as the sole carbon and nitrogen sources (Table 2). The protease production by B. stearothermophilus F1 was regulated by the concentration of peptone iv. Peptone at 1% gave the best yield but further increases in the concentration of peptone iv reduced the protease production due to the accumulation of nitrogenous catabolites such as amino acids and ammonia. Growth and protease production by B. stearothermophilus F1 was also detected in non-protein medium although at lower growth rates. Protease production in sodium nitrate medium was only about 11% of that with peptone iv, while ammonium compound completely inhibited the production. Apparently, the used of amino acids as nitrogen source had repressed and reduced protease production and the growth of B. stearothermophilus F1. Table 2: Effect of nitrogen source on protease production. Nitrogen Source (1%) Peptone iv Soytone Corn steep liqour Gelatine Casein Beef extract Peptone I Polypeptone Tryptone Yeast extract Urea Casamino acid Sodium nitrate Ammonium sulfate Ammonium nitrate Asparagine Aspartic Leucine Cysteine Arginine Glycine (Source: Rahman et al., 2003)

Protease Activity (U/ml) 9520 ± 115 6560 ± 17 1022 ± 13 860 ± 40 730 ± 62 690 ± 40 0 0 0 0 50 ± 10 0 1963 ± 47 45 ± 13 60 ± 10 433 ± 61 217 ± 76 0 0 0 0

Growth (OD 679) 0.70 ± 0.01 0.68 ± 0.03 0.60 ± 0.05 0.55 ± 0.01 0.48 ± 0.04 0.58 ± 0.02 0.56 ± 0.07 0.60 ± 0.01 0.58 ± 0.03 0.50 ± 0.01 0.20 ± 0.07 0.10 ± 0.01 0.45 ± 0.05 0.29 ± 0.08 0.23 ± 0.09 0.20 ± 0.06 0.28 ± 0.08 0.15 ± 0.05 0.10 ± 0.03 0.18 ± 0.04 0.10 ± 0.07

The presence of additional carbon source in peptone iv medium had resulted in the reduction of protease production by B. stearothermophilus F1, with the exception of raffinose (Figure 4). The addition of these carbon sources, however, slightly increased the growth by B. stearothermophilus F1 in the range of 0.68 to 0.82. Addition of raffinose to the protein medium slightly increased the protease production to 6%, while the addition of galactose slightly decreased its production to 5% (Rahman et al., 2003). Low level of activity was obtained with starch, sorbitol, trehalose, sucrose and mannose. On the other hand, addition of

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most carbon sources in sodium nitrate medium inhibited its production with an exception for meliobiose and trehalose (Figure 5).

Figure 4: Effect of additional carbon source on protease production in peptone iv medium.

Calcium was not required for protease formulation but its addition in the basal medium enhanced the enzyme yield. Addition of 4.5mM calcium maximized protease production by stabilizing the secreted enzyme, but a higher concentration lowered the enzyme production. Strontium was equally effective as an inducer for enzyme production, while Mn2+ and Fe2+ significantly inhibited the enzyme production but little effect on the growth.

Figure 5: Effect of carbon source on protease production in sodium nitrate medium.

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Recombinant Protease A simple method for obtaining enzymes with improved thermostability is to isolate the enzyme from naturally occurring thermophilic organisms. However, the main disadvantage of this approach is that it is not practical to produce large quantities of enzymes from such organisms, as the yield is normally low because the enzymes tend to be susceptible to denaturation upon longer incubation periods. Therefore, gene encoding F1 serine protease was cloned in prokaryotic system (Hamid, 2000). An attempt has been made to co-express the F1 protease with bacteriocin release protein (BRP) to form permeable zones for secretory expression. The recombinant F1 protease was efficiently secreted into the culture medium using E. coli XL1-Blue harbouring two vectors: pTrcHis bearing the F1 protease gene and pJL3 containing BRP gene. Both vectors contain the Escherichia coli lac promoter-operator system. In the presence of 40µM IPTG, the recombinant F1 protease and the BRP were expressed and mature F1 protease was released into the culture medium (Fu et al., 2003). Analysis of SDS-PAGE shown in Figure 6 revealed that F1 protease was present in the culture medium, but it was not the only protein. However, a comparison of the proteins intensity demonstrated that F1 protease was relatively enriched in the medium with a higher amount of F1 protease after 72 h incubation (lane 8). The respective band was further confirmed with native SDS-PAGE through activity staining.

Figure 6: SDS–PAGE analysis of F1 protease excretion by the E. coli XL1-Blue cells harboring the two plasmids (pJL3 and recombinant pTrcHis). Note: M, standard marker. Lane 1, the total cell lysate from 24 h cultivation. Lane 2, the total cell lysate from 30 h cultivation. Lane 3, the total cell lysate from 48 h cultivation. Lane 4, the total cell lysate from 72 h cultivation. Lane 5, the medium concentrated from 24 h cultivation cells. Lane 6, the medium concentrated from 30 h cultivation cells. Lane 7, the medium concentrated from 48 h cultivation cells. Lane 8, the medium concentrated from 72 h cultivation cells. The E. coli cells were grown in LB broth in the presence of 40µM IPTG. Arrow indicates the F1 protease. (Source: Fu et al., 2003)

The wild-type F1 protease was purified to homogeneity by heat treatment, ultrafiltration and gel filtration chromatography (Rahman et al., 1994). Heat treatment (3 h, 70°C) removed 40% of the protein while retaining 100% of the initial protease activity (Table 3). The enzyme fraction was eluted as a single peak of proteolytic activity on Sephadex G-100 column after concentrating the enzyme through ultrafiltration. The final purification step of purification resulted in removal of more than 99% of crude protein with a good recovery. A 128-fold purification at 75% high recovery was obtained due to the enzyme stability during purification steps. Meanwhile, the recombinant F1 protease was purified by a one-step

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procedure following the concentration of the original crude enzymes by freeze-drying, which was then heat-treated at 70°C (Fu et al., 2003). The heat-treatment method provides a simple and effective method for purifying the recombinant F1 protease to near homogeneity with no activity lost. The specific activity of the enzyme was also increased four fold after this step (Table 3). The purified enzyme was checked for purity by SDS-PAGE. Gels were stained with Coomassie blue R-250, and one major band was visible, which migrated at approximately 27 kDa (Figure 7). Table 3: Purification of the native and recombinant thermostable F1 protease

Purification Steps

Total Total protein Specific activity activity(U) (mg) (U/mg)

Recovery (%)

Purification fold

Native F1 protease Crude Heat treated Ultrafiltration G-100 G-100

294,800 294,000 294,500 260,800 223,300

101.4 62.4 19.24 0.8325 0.599

2909 4712 15307 313273 372963

100 100 99 88 75

1.0 1.6 5.3 108.0 128.3

Recombinant F1 protease Crude Heat treated

11,000 10,000

24.6 6.1

447 1790

100 99

1.0 4.0

Characterization of F1 Protease Effect of pH on Enzyme Activity and Stability The effect of pH ranging from 5.0 to 12.0 at 70°C on the native and recombinant F1 protease activity was investigated. The maximum activity for the azocasein hydrolysis was observed at pH 9.0. The native F1 protease had 50% of its maximum activity at pH 5.0 and 11.0 respectively (Rahman et al., 1994). However, the recombinant F1 protease showed only minor activity when tested at pH 5.0, and less than 30% of its maximum activity was obtained at pH 12.0, while more than 50% of its maximum activity was retained at pH 11.0 (Fu et al., 2003). The effect of pH on recombinant F1 protease stability was examined in the pH range between 5.0-12.0. Rahman et al. (1994) had reported that the native F1 protease was stable between pH 6.0 to 11.0 when incubated for 30 min at 70°C. Meanwhile, the recombinant F1 protease was stable between pH 5.0 to 12.0 when incubated for 30 min at 70°C (Fu et al., 2003). However, F1 protease was found to be stable between pH 8.0 to 10.0, when the enzyme was further incubated for 24 h at 70°C (Rahman et al., 1994; Fu et al., 2003).

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Effect of Temperature on Protease Activity and Stability The native and recombinant F1 protease were incubated at different temperatures ranging from 50°C to 90°C in the presence of 2 mM CaCl2. The optimal temperature for recombinant F1 protease activity was around 80°C. Although the activity of recombinant F1 protease at 90°C was higher than that at 50°C but less than 50% of recombinant protease activity was retained.

Figure 7: SDS–PAGE (12%, w/v) of the purified protease. Note: Lane M, molecular mass markers (in kDa). Lane 1, crude enzyme. Lane 2, purified enzyme by heat-treatment. Arrow indicates the recombinant F1 protease. (Source: Fu et al., 2003)

On the other hand, the native F1 protease had the optimal temperature for the protease activity around 85°C and retained higher activity at 90°C as compared to recombinant F1 protease (Rahman et al. 1994). However, this property was not retained when the F1 protease was expressed in E. coli system. The recombinant enzyme might be improperly folded and/or assembled at 37°C while the native enzyme was produced at a higher growth temperature (65°C). However, the optimum temperatures for both forms of F1 proteases were higher than that of thermolysin (80°C) and subtilisin (60°C). Both the recombinant and native F1 protease showed similar half-lives of 4 h at 85°C and 25 min at 90°C (Salleh et al., 1997; Fu et al., 2003). The native F1 protease was stable for 9 h at 80°C without significant loss of enzyme activity and was far more stable than thermolysin and subtilisin which were completely denatured within 4 h and 1 h, respectively (Salleh et al., 1997).

Substrate Specificity The protease was active on a variety of modified (azocasein and azocoll) and natural proteins (bovine serum albumin, casein and haemoglobin) substrates. The protease exhibited

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the highest activity with casein. Results with sythethic substrates confirmed that no trypsinlike activity was present as a chromogenic trypsin substrate, BAPNA was not hydrolyzed when tested. This was further supported as none of the ester substrates of trypsin (BAEE, BAME and TAME) and ester substrates of chymotrypsin (ATEE and BTEE) tested were hydrolysed by the enzyme. Therefore, judging from the effect of inhibitor and substrate specificity studies, it can be concluded that the enzyme did not exhibit trypsin-like activity. F1 protease exhibited specificities towards hydrophobic amino acid residues such as phenylalanine at the carboxyl side of the splitting point (Rahman et al., 1994).

Effect of Inhibitors The effect of various classes of inhibitors on the native and recombinant F1 protease activities was investigated. Inhibitors of sulfhadryl protease, indoacetamide (IAA); metalloprotease, EDTA; aspartic protease, pepstatin and aminopeptidase, bestatin did not inhibit the F1 protease activity. However, the protease activity was totally inhibited by the serine protease inhibitor, phenylmethanesulphonyl flouride (PMSF) at 10 mM. The results clearly indicate that this enzyme belongs to the serine protease family and it is comparable to the native protease from B. stearothermophilus F1 (Rahman et al., 1994; Fu et al., 2003).

Figure 8: The effect of calcium (Ca2+) and manganese (Mn2+) on the thermostability of protease at 95°C. The enzyme was preincubated with different concentration of Ca2+ ( ) and Mn2+ ( ) for 5 min at 95°C. (Source: Rahman et al., 1994)

Effect of Ca2+ and Mn2+ on Thermostability Most metal ions tested did not greatly affect the enzyme activity. However, Ca2+ (slightly) and Mn2+ (strongly) enhanced its activity. Manganese at 10 mM activated the protease activity by two fold. However, the thermostability conferred by Ca2+ was much greater than that of Mn2+ (Figure 8). The residual activity at 95°C increased from about 50%

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in 2 mM Ca2+ to 70% in 14 mM Ca2+. On the other hand, the additions of Mn2+ even up to 20 mM only increase the residual activity about 10%. Even though both Ca2+ and Mn2+ activated the F1 protease activity, apparently only Ca2+ satisfied the structural requirement for stabilization at higher temperatures (Rahman et al., 1994).

REFERENCES Adinarayana, K., Elaiah, P., and Prasad, D. S., 2003. Purification and partial characterization of thermostable serine alkaline protease from newly isolated Bacillus substillis PE-11. AAPS PharmSciTech, 4(56): 1-9. Banarjee, U. C., Sani, R. K., Azmi, W., and Soni, R., 1999. Thermostable alkaline protease from Bacillus brevis and its characterization as a laundry detergent additive. Process Biochemistry, 35: 213- 219. Brown, C. M., Campbell, I., and Priest, F. G., 1987. Introduction to Biotechnology, Backwell Scientific Publication, Oxford. pp: 51-52. Choi, I. G., Barg, W.G., Kim, S.H., and Yu, Y.G., 1999. Extremely thermostable serine-type protease from the Aquifex pyrophilus. Journal of Biological Chemistry, 274(2): 881-888. Cowan, D. A., and Daniel, R. M., 1982. Purification and some properties of extracellular protease (caldolysin) from an extreme thermophile. Biochimica et Biophysica Acta, 705: 293-305. Desai, A. J., and Dhala, S. A., 1969. Purification and properties of proteolytic enzymes from thermophilic actinomycetes. Journal of Bacteriology, 100(1): 149-155. Durham, D. R., Stewart, D. B., and Stellwagt, E. J., 1987. Novel alkaline and heat-stable serine proteases from alkalophilic Bacillus sp. strain GX6638. Journal of Bacteriology, 169(6): 2762-2768. Fu, Z., Hamid, S., Razak, C. N., Basri, M., Salleh, A. B., and Rahman, R. N. Z. A., 2003. Secretory expression in Escherichia coli and single-step purification of a heat-stable alkaline protease. Protein Expression and Purification, 28(1): 63-68. Fujiwara, N., and Yamamoto, K., 1987. Production of alkaline protease in low-cost medium by alkalophilic Bacillus sp. and properties of the enzyme. Journal of Fermentation Technology, 65(3): 345-348. Fujiwara, N., Masui, A., and Imanaka, A., 1993. Purification and properties of the highly thermostable alkaline protease from an alkaliphilic and thermophilic Bacillus sp. Journal of Biotechnology, 30(2): 245-256. Fusek, M., Lin, X. L., and Tang, J., 1990. Enzymic properties of thermopsin. Journal of Biological Chemistry, 265(3): 1496-1501 Gessesse, A., Hatti-Kaul, R., Gashe, B. A., and Mattiasson, B., 2003. Novel alkaline proteases from alkaliphilic bacteria grown using chicken feather. Enzyme and Microbial Technology, 32(5): 519-524. Godfrey, T., and West, S., 1996. Introduction to industrial enzymology. In: Industrial Enzymology, 2nd Ed., (eds.) Godfrey, T. and West, S., Macmillan Publisher Inc., New York. pp: 1-8. Gusek, T., and Kinsella, J. E., 1987. Purification and characterization of the heat-stable serine proteinase fom Thermomonospora fusca YX. Biochemical Journal, 246(2): 511-517.

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Hamid, A. S., 2000. Cloning and expression of alkaline protease gene from Bacillus stearothermophilus strain F1. M. S. Thesis, Universiti Putra Malaysia, Serdang, Malaysia. Kannan, Y., Koga, Y., Inoue, Y., Haruki, M., Takagi, M., Imanaka, T., Morikawa, M., and Kanaya, S., 2001. Active subtilisin-like protease from a hyperthermophilic archaeon in a form with a putative prosequence. Applied and Environmental Microbiology, 67(6): 2445- 2452. Klingeberg, M., Hashwa, F., and Antranikian, G., 1991. Properties of extremely thermostable proteases from anaerobic hyperthermophilic bacteria. Applied Microbiology and Biotechnology, 34(6): 715-719. Kubo, M., Murayama, K., Seto, K., and Imanaka, T., 1988. Highly thermostable neutral protease from Bacillus stearothermophilus. Journal of Fermentation Technology, 66(1):13-17. Lin, X., and Tang J., 1990. Purification, characterization and gene cloning of thermopsin, a thermostable acid protease from Sulfolobus acidocaldarius. Journal of Biological Chemistry, 265(3): 1490- 1495. MacIver, B., Michale, R. H., Saul, D. J., and Bergquist, P. L., 1994. Cloning and sequencing of a serine protease gene from a thermophilic Bacillus species and its expression in Escherichia coli. Applied and Environmental Microbiology, 60(11): 3981-3988. Manachini, P. L., Fortina, M. G., and Parini, C., 1988. Thermostable alkaline protease produced by Bacillus thermoruber - a new species of Bacillus. Applied Microbiology and Biotechnology, 28(4): 409-413. Matsuzawa, H., Takugawa, K., Hamaoki, M., Mizoguchi, M., Taguchi, H., Terada, A., Kwon, S.T., and Ohta T., 1988. Purification and characterization of aqualysin 1, (a thermophilic alkaline serine protease) produced by Thermus aquaticus YT-1. European Journal of Biochemistry, 171(3): 441-447. Matsuzawa, H., Hamaoki, M., and Takanida, O., 1983. Production of thermophilic extracellular proteases (aqualysin 1 and 11) by Thermus aquaticus YT-1, an extreme thermophiles. Agricultural Biological Chemistry, 47: 25-28. Meloun, B., Baudys, M., Kostka, V., Hausdorf, G., Frömmel, C., and Höhne, W.E., 1985. Complete primary structure of thermitase from Thermoactinomyces vulgaris and its structural features related to the subtilisin-type proteinase. FEBS Letters, 183(2): 195200. Ohta, Y., Ogura, Y., and Wada, A., 1966. Thermostable protease from thermophilic bacteria. 1. Thermostability, physicochemical properties and amino acid composition. Journal of Biological Chemistry, 241(24): 5919-5925. Olędzka, G., Dąbrowski, S., and Kur, J., 2003. High-level expression, secretion, and purification of the thermostable aqualysin I from Thermus aquaticus YT-1 in Pichia pastoris. Protein Expression and Purification, 29(2): 223-229. Patel, T. R, Jackman, D. M., and Bartlett, F. M., 1983. Heat-stable protease from Pseudomonas fluorescens T16: Purification by affinity column chromatography and characterization. Applied and Environmental Microbiology, 46(2): 333-337. Peek, K., Daniel, R. M., Monk, C., Parker, L., and Coolber, T., 1992. Purification and characterization of a thermostable proteinase isolated from Thermus sp. strain Rt41A. European Journal of Biochemistry, 207: 1035-1044.

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Peek, K., Veitch, D. P., Prescott, M., Daniel, R. M., MacIver, B., and Bergquist, P. L., 1993. Some characteristics of a proteinase from a thermophilic Bacillus sp. expressed in Escherichia coli: comparison with the native enzyme and its processing in E.coli and in vitro. Applied and Environmental Microbiology, 59(4): 1168-1175. Rahman, R. N. Z. A., Basri, M., and Salleh, A. B., 2003. Thermostable alkaline protease from Bacillus stearothermophilus F1; nutritional factors affecting protease production. Annals of Microbiology, 53(2):199-210. Rahman, R. N. Z. A., Razak, C. N., Ampon, K., Basri, M., Yunus, W. M. Z., and Salleh, A. B., 1994. Purification and characterization of a heat-stable alkaline protease from B. stearothermophilus F1. Applied Microbiology and Biotechnology, 40(6): 822-827. Rao, M. B., Tanksale, A. M., Ghatge, M. S., and Deshpande, V. V., 1998. Molecular and biotechnological aspects of microbial proteases. Microbiology and Molecular Biology Reviews, 62(3): 597-635. Razak, C. N., Rahman, R. N. Z. A., Ampon, K., Basri, M., Yunus, W. M. Z., and Salleh, A. B., 1995. Production of a thermostable alkaline serine protease by a new strain of Bacillus stearothermophilus. Journal of Bioscience, 6(1): 94-100. Salleh, A. B., Rahman, R. N. Z. A., Basri, M., and Razak, C. N. A. 1997. The effect of temperature on the protease from Bacillus stearothermophilus strain F1. Malaysian Journal of Biochemistry and Molecular Biology, 2: 37-41. Saravani, G. A., Cowan, D. A., Daniel, R. M., and Morgan, H. M., 1989. Caldolase, a chelator-insensitive extracellular serine proteinase from Thermus sp. Biochemical Journal, 262(2): 409-416. Saul, D. J., Williams, L. C., Toogood, H. S., Daniel, R. M., and Bergquist, P. L., 1996. Sequence of the gene encoding a highly thermostable neutral proteinase from Bacillus sp. strain EA1: expression in Escherichia coli and characterisation. Biochimica et Biophysica Acta, 1308(1): 74-80. Schalk, C., Remy, J. M., Chevrier, B., Moras, D., and Tarnus, C., 1992. Rapid purification of the Aeromonas proteolytica aminopeptidase: crystallization and preliminary X-Ray data. Archives of Biochemistry and Biophysics, 294: 91-97. Sidler, W., and Zuber, H., 1972. Neutral proteases with different thermostabilities from a falcutative strain of Bacillus stearothermophilus grown at 40°C and 50°C. FEBS Letters, 25(2): 292-294. Sidler, W., and Zuber, H., 1977. The production of extracellular thermostable neutral proteinase and α-amylase by Bacillus stearothermophilus. Applied Microbiology and Biotechnology, 4(4): 255-266. Stepaniak, L., Fox, P.F., and Daly, C., 1982. Isolation and general characterization of a heatstable proteinase from Pseudomonas fluorescens AFT 36. Biochimica et Biophysica Acta, 717(2): 376-383. Suzuki, Y., Ito, N., Yuki, T., Yamagata, H., and Udaka, S., 1989. Amino acid residues stabilizing a Bacillus α–amylase against irreversible thermoinactivation. Journal of Biological Chemistry, 264(32): 18933-18938. Takami, H., Akiba, T., and Horikoshi, K., 1989. Production of extremely thermostable alkaline protease from Bacillus sp. no AH-101. Applied Microbiology and Biotechnology, 30(2): 120-124.

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Tsuboi, R., Sanada, T., Takamori, K., and Ogawa, H., 1987. Isolation and properties of extracellular proteinases from Sporothrix schenckii. Journal of Bacteriology, 169(9): 4104-4109. Van Noort, J. M., Van den Berg, P., and Mattern, I. E., 1991. Visualization of proteases within a complex sample following their selective retention on immobilized bacitracin, a peptide antibiotic. Analytical Biochemistry, 198(2): 385-390. Voorhorst, W. G. B., Warner, A., de Vos, W. M., and Siezen, R. J., 1997. Homology modelling of two subtilisin-like serine proteases from the hyperthermophilic archaea Pyrococcus furiosus and Thermococcus stetteri. Protein Engineering, 10(8): 905-914. Wu, J., Bian, Y., Tang, B., Chen, X., Shen, P., and Peng, Z., 2004. Cloning and analysis of WF146 protease, a novel thermophilic subtilisin-like protease with four inserted surface loops. FEMS Microbiology Letters, 230(2): 251- 258.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 95-110

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 6

ORGANIC SOLVENT TOLERANT PROTEASES Azira Muhammad, Raja Noor Zaliha Raja Abd. Rahman, Abu Bakar Salleh and Mahiran Basri Universiti Putra Malaysia, Malaysia

ABSTRACT The potential advantage of using organic solvents for enzymatic reactions was due to the shift of reaction equilibrium of hydrolytic enzymes towards completion of the synthetic reaction. However, the use of these solvents normally led to loss of most enzyme activities. Therefore, proteases, which were naturally stable in the presence of organic solvents, were very useful for synthetic reactions. In our laboratory, several organic solvent-tolerant protease producers have been successfully isolated from benzene–toluene–xylene–ethylbenzene (BTEX) tolerant bacteria. The formulation of physical and nutritional factors affecting the enzymes production have led to the optimized production of proteases from both Pseudomonas aeruginosa strain K and Bacillus cereus 146, which were stable in organic solvents with Po/w values between 4.0 and 8.8. Due to these remarkable properties, strain K protease with a molecular size of 51.0 kDa was further purified by two purification steps and characterized. Complete nucleotide sequences from Pseudomonas aeruginosa strain K and Bacillus pumilus 115b were also obtained and analyzed to gain better understanding of its nucleotides which responsible for the enzymes biocatalytic interaction with solvent environment.

INTRODUCTION Proteases are useful for applications in the production of useful products by peptide synthesis in organic media. However, the process is limited due to the specificity and the instability of this enzyme in the presence of organic media. Therefore, the discovery of proteases active in organic solvents has greatly expanded their potential in the syntheses of useful products.

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The advantages of using organic solvents as the media for enzymatic reactions are due to the shift in the reaction equilibrium of hydrolytic enzymes towards the completion of synthetic reaction, the possibility to eliminate undesirable side reactions of inorganic materials in the media and the substrates besides avoiding product inhibitions. Microbial proteases that can function as catalyst in non-aqueous solvents presents new prospects, such as increasing the solubility of hydrophobic substrates and controlling the specificity by improving thermal stability of enzymes. Further interest stems from the fact that many hydrophobic solvents are classified as environmental pollutants and the degradation of these solvents by microorganisms is the focus of the bioremediation industry (Cruden et al., 1992). It was found that solvent-tolerant microorganisms were also useful in biotransformations with the cells in two-phase solvent-water system. Hence, the search for proteases that are naturally stable in the presence of organic solvents has become an essential area in enzymology. The addition of organic solvent to the reaction mixture will reduce molar fraction of water. When the polarity of a medium surrounding enzyme molecules is reduced, the hydrophobic domains are apt to disperse, resulting in the unfolding of the molecules (Tanaka and Kawamoto, 1991). In the absence of bulk water, their activities can be modulated, their selectivities tailored and their stabilities altered (Klibanov, 2001). However, the main issue concerning nonaqueous enzymology is often the drastic drop in enzymatic activity in organic solvents. Studies have been carried out to explain the causes of this phenomenon and many methods have been devised in order to enhance the enzymatic activity in organic solvents. Protein conformation in organic media is critical to elucidate our understanding on the mechanisms of the enzyme. When enzymes are placed in organic solvents containing little or no water, they exhibit unusual properties, including the ability to catalyze additional reactions, markedly altered their specificity and “molecular memory” (Fitzpatrick et al., 1993). Subtilisin Carlsberg has been extensively studied to understand the basis of these phenomena. The crystal structure of the serine protease subtilisin Carlsberg in anhydrous acetonitrile was determined at 2.3Å resolution (Fitzpatrick et al., 1993). It was found that the structure was essentially identical to the three-dimensional structure of the enzyme in water, where one-third of all subtilisin-bound acetonitrile molecules reside in the active center, occupying the same region (P1, P2 and P3 binding sites) as the specific protein inhibitor eglin c. This observation is consistent with the preference of subtilisin for substrates having a polar backbone and hydrophobic side chains. Further insight into the enzyme mechanism in organic solvents was provided by an x-ray crystal structure of an acyl-enzyme intermediate of subtilisin Carlsberg formed in acetonitrile (Schmitke et al., 1998). This structure was compared to the enzyme structure in water and it was found that the acyl-enzyme covalent intermediate of the two structures were virtually identical. Thus it was concluded that acylation in either aqueous or nonaqueous solvents caused no appreciable conformational changes. These findings verified kinetic evidence that the enzyme mechanisms in organic solvents and in water are similar. However, a distinct binding of the solvent molecules in the active site between the acyl-enzyme intermediate and the free enzyme structures in different solvents might be responsible for at least some of the heretofore unexplained disparity in the activity of enzymes in organic solvents. Other protease structures in neat organic solvents that have been determined include γ-chymotrypsin in hexane (Yennawar et al., 1994) and elastase in acetonitrile (Allen et al., 1996). Structural studies that have been carried out on several proteases have given further insights into the question of why enzymatic activity is often much reduced in organic solvents

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compared to water. The information gathered from structural and mechanistics studies have helped in the use of proteases for the synthesis of peptides, esters and other biologically active organic molecules where many efforts have been devoted to find conditions where enzymes can be used in combination with organic solvents while obtaining their enzymatic activity.

SOURCES OF ORGANIC SOLVENT-TOLERANT PROTEASE Little attention has been given to the isolation and the study of microorganisms that produce organic solvent-stable proteases. Natural proteases which are organic solvent-stable have been reported from solvent-tolerant Pseudomonas sp. (Ogino et al., 1995; Gupta et al., 2005) and Bacillus sp. (Ghorbel et al., 2003). An organic solvent-tolerant Pseudomonas aeruginosa PST-01, which secreted an organic solvent-stable protease, was isolated from soil following a two-step procedure: high proteolytic enzyme producers were first isolated, and then the organic solvent-tolerant microorganism was selected from these high-rate proteolytic enzyme producers (Ogino et al., 1995). The stability of the enzyme in the presence of organic solvents of which the values of the log P were equal to or more than 3.2, was almost the same as that in the absence of organic solvents. Another solvent-stable protease from the same microorganism was reported by Gupta et al., (2005) where solvent-tolerant P. aeruginosa PseA strain was isolated from soil by cyclohexane enrichment. This strain secreted considerable amount of extracellular solventstable protease with higher specific activity as compared to other solvent-tolerant Pseudomonas proteases such as PST-01. Besides protease-producing Pseudomonas species, an organic solvent-tolerant Bacillus cereus producing organic solvent-stable protease was isolated from a fishing industry wastewater (Ghorbel et al., 2003). The strain that showed high-activity alkaline protease was identified as B. cereus BG1. Stability studies of protease from B. cereus BG1 were carried out and the enzyme was found to retain more than 95% of its initial activity in the presence of 25% methanol, DMSO, acetonitrile and DMF. The optimum temperature for protease activity was recorded at 60ºC in the presence of 2mM Ca2+ and 50ºC in the absence of Ca2+. At 60°C, Ca2+ stimulated the protease activity by 500%. The optimum pH was 8.0 and the enzyme was quite stable in various pH buffers between pH 6.0 and 9.0 when incubated at 50ºC for 1 and 3h. Enzyme activity was inhibited by EDTA, suggesting a metalloprotease enzyme (Ghorbel et al., 2003).

PURIFICATION AND CHARACTERIZATION OF ORGANIC SOLVENT-TOLERANT PROTEASE PST-01 protease was purified by successive hydrophobic interaction chromatography using Butyl-Toyopearl gels. The purified protease had a molecular mass of 38 kDa and the optimum temperature and pH for casein hydrolysis were 55ºC and 8.5, respectively (Ogino et al., 1999). PST-01 protease was stable at pH 8-12 and below 50ºC and was determined to be a metalloprotease which was inhibited by EDTA,1,10-phenantroline and phosphoramidon. In

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general, PST-01 protease was more stable than commercially available proteases, namely, subtilisin Carlsberg, thermolysin and α-chymotrypsin (Ogino et al., 1999). A study on the effect of disulfide bonds present on PST-01 protease showed that the bond between Cys-30 and Cys-58 played an important role in not only the heat stability but also the organic solvent stability (Ogino et al., 2001). Solvent-stable protease from Pseudomonas aeruginosa PseA was purified by combination of ion exchange and hydrophobic interaction chromatography using QSepharose and Phenyl Sepharose 6 Fast Flow matrix, respectively. The apparent molecular mass based on SDS-PAGE was estimated to be 35 kDa with 8.0 as the optimum pH. The enzyme was stable in the pH range of 6.0-9.0. The protease was most active at 60ºC and was characterized as metalloprotease because of its sensitivity to EDTA and 1,10-phenantroline. PseA protease withstands a range of detergents, surfactants and solvents. It is stable and active in all of the solvents having log P above 3.2, at least up to 72h (Gupta et al., 2005).

OUR WORK Isolation and Screening of Organic Solvent-tolerant Protease An extracellular organic solvent-tolerant protease producer has been successfully isolated from an organic solvent-tolerant microorganism and identified as Pseudomonas aeruginosa strain K (Geok et al., 2003). The bacteria were isolated from contaminated soils of a wood factory in Selangor, Malaysia. From the screening on skim milk agar (SMA), eleven bacteria which showed positive results by forming zones of lysis around the colonies (Figure 1) on SMA were isolated and purified. These isolates were reported to be benzene-toluene-xyleneethyl benzene (BTEX) tolerant bacteria. On the basis of the relative stability in 25% (v/v) benzene and toluene, strain K was selected as the most potent producer of organic solventtolerant proteolytic enzyme.

Figure 1: Zones of Lysis on SMA Plate. Pseudomonas aeruginosa strain K was grown and screened qualitatively for the protease production on skim milk agar containing (g/L): skim milk powder, 12.0 and nutrient agar, 13.8. The plate was incubated at 37°C for 24 h. Proteolytic enzyme produced by strain K hydrolysed the skim milk and formed clearing zones around the colonies on skim milk agar plate (Source: Geok, 2003).

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Another organic solvent-tolerant protease producer was isolated from bacterium designated as 146. The strain was a Gram-positive, spore-forming, nitrate-positive, rodshaped organism capable of hydrolizing gelatin, starch, skim milk and identified as Bacillus cereus. This strain was shown to be BTEX-tolerant bacteria and proven to be polycyclic aromatic hydrocarbons (PAHs) degrader (Shafee et al., 2006). An organic solvent-tolerant protease producer was isolated from an organic solventtolerant microorganism identified as B. pumilus 115b (Mahamad, 2006). The bacterium was screened from 5 isolates of BTEX-tolerant bacteria. Isolate 115b was found to secrete the highest protease production and was highly stable in 25% (v/v) benzene and toluene.

Physical Factors Affecting Growth and Protease Production Physical factors affecting the production of solvent-tolerant protease strain K was investigated (Rahman et al., 2005a). Investigations on the effect of cultivation temperatures on cell growth and protease production at different incubation temperatures: 4, 37, 40, 45, 50 and 60°C have been carried out at OD540, nutrient agar plates and basal media, respectively. Growth and protease production were detected from 37 to 45ºC with 37ºC being the optimum temperature. Higher temperatures at 40°C and 45°C caused more than 24.0% and 53.0% lost of the enzyme activity at 48h incubation. No bacterial growth and protease activity were observed at 4°C or at 50°C or higher temperatures. Hence, based on the optimum temperature, Pseudomonas aeruginosa strain K was classified as mesophiles. The amount of inoculum used to culture the bacteria can affect the protease production of Pseudomonas aeruginosa strain K. The result indicated that, protease production and bacterial growth were optimum when 4.0% (v/v) of bacterial inoculum was used and the culture was incubated for 48 h (Table 1). Higher inoculum size at 6.0% (v/v), 8.0% (v/v) and 10.0% (v/v) decreased the protease production by 28.2%, 54.9% and 60.6%, respectively. A drop of optical density (O.D540) from O.D 1.130 to 1.003 was observed in bacterial growth if the inoculum size was increased. Inoculum size 8.0% (v/v) and 10.0% (v/v) also decreased 13.5% and 15.8% of the bacterial growth compared to the control, respectively (Table 1). Therefore, high inoculum sizes might not necessarily give higher protease yield or cell growth. On the contrary, higher inoculum sizes could result in the lack of oxygen and nutrient depletion in the culture media. Agitation rates influenced the protease yield and cell growth of strain K. Maximum protease production and bacterial growth were observed when the culture medium was grown under static condition. Maximum enzyme activity was achieved at static conditions with 4.0% (v/v) inoculum while protease production was decreased by shifting the culture from stationary to shaking condition. At acidic pH (pH 6.0), 63.2% reduction of the enzyme was observed compared to pH 7.0. At pH 8.0, the protease yield was decreased by 16.9% after 48 h incubation compared to pH 7.0 (Table 2). Extracellular protease was detected over a broad pH range (6.0 to 9.0), with optimum production exhibited at pH 7.0 (Table 2).

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Nutritional Factors Affecting Growth and Protease Production Nutritional requirements for optimized production of organic solvent-tolerant strain K protease were investigated (Rahman et al., 2005b). Media containing lactose, fructose and arabinose showed enhancement of protease activity by as much as 261%, 162% and 214%, respectively. Maximum protease activity was achieved with sorbitol as the sole carbon source, giving 362% increase in protease yield compared to the basal medium, followed by starch and lactose at pH 7.0 and 37ºC (Figure 2). Table 1: Effect of Inoculum Sizes on Strain K Protease Production Relative activity Inoculum size (%) 2 4 6 8 10

24h 0.20 1.00 0.36 0.35 0.36

48h 0.49 1.00 0.72 0.45 0.39

Note: Culture media were inoculated with 2.0, 4.0, 6.0, 8.0, 10.0 % (v/v) of inoculum and incubated at 37°C with shaking at 150 rpm for 24 h and 48 h . Protease activity at 4.0% (v/v) inoculum was taken as 1.0.

Table 2: Effect of pH on Strain K Protease Production Relative activity pH 5 6 7 8 9 10 11

24h 0 0.21 1.00 0.08 0.36 0 0

48h 0 0.37 1.00 0.83 0 0 0

Note: Bacterial cultures were adjusted to pH 5.0, 6.0, 7.0, 8.0, 9.0, 10.0, 11.0 and incubated at 37°C for 24h and 48h. Protease activity at pH 7.0 was taken as 1.0.

The best organic nitrogen source was casamino acid which produced 425 % increase in relative activity after 48 h incubation. Tryptone, soytone and yeast extract supported protease production while corn steep liquor and beef extract inhibited the protease activity. Protease production was detected in medium containing a non-organic nitrogen source. Significant protease production was observed with sodium nitrate as a sole nitrogen source however, ammonium nitrate completely inhibit it. The possible synergistic effect produced by combining both the organic and inorganic nitrogen sources was also examined. The best

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combination for protease activity reported was casamino acids (1.0 % w/v) and sodium nitrate (0.1 % w/v) which give a 35% increase in protease and 43% increase in cell growth. The effect of amino acids on protease production was studied and it was found that more than 62% drop in production occurred in the presence of glutamine and histidine after 48h incubation. The same amino acids reduced bacterial growth by 62.5% and 75% respectively. Moreover, cysteine was found to cause a complete inhibition of protease production even though casamino acids and sodium nitrate were present in the basal medium.

4

Realative activity

3.5 3 2.5 2 1.5 1 0.5

G G lu co M se al to Fr se uc to La se ct Ar os ab e in o So se rb ito St l ar De ch xt r M ose an no Su se cr os e

M

+ M

P +

M M

M

M

BM

0

Carbon source (1.0% w/v) Figure 2: Effect of Carbon Sources on Strain K Protease Production. Different carbon sources: 1.0% (w/v) of lactose, sucrose, maltose, fructose, arabinose, sorbitol, dextrose, mannose, starch were added into minimal media with peptone, type iv (MM + P) and incubated at pH 7.0, 37°C for 24h ( ) and 48h ( ). BM, basal medium; MM + P, minimal medium with the presence of peptone, type iv; MM, minimal medium; MM + G, minimal medium with the presence of glycerol. The relative activity was based on protease activity in the culture media relative to the basal medium. (Source: Rahman et al., 2005b)

Metal ions were required in the fermentation media for optimum production of proteases. The enzyme production was maximized with the addition of metal ions such as K+, Mg2+ and Ca2+. However, the presence of Fe3+, Mn2+ and Li+ decreased proteolytic activity by more than 86% after 48 h incubation. Nutritional factors affecting organic solvent-tolerant protease production by a Bacillus cereus strain 146 was investigated (Shafee et al., 2006; Shafee et al., 2005). The bacterium was shown to require lactose as a carbon source and peptone as a nitrogen source. The optimum fermentation condition for production of the protease was in the presence of glucose and beef extract (Shafee et al., 2005). Optimum pH was determined to be at 10.0 at incubation temperature of 37°C for 48h. At this temperature, protease activity was at 0.92 µg/ml/min, an increase of approximately 46% from incubation time of 24h (Figure 3).

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1

Relative activity

0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0 25

30

37

40

45

Temperature (o C) Figure 3: Effects of temperature on protease production and growth of B. cereus 146. Bacterial cultures in media M4 (Singh et al., 2000) were incubated at different incubation temperatures at 150 rev/min agitation rate. Protease activity was determined following 24 ( ), 48 ( ) and 72 h ( ) of incubation. Results are means of three independent determinations (Source: Shafee et al., 2006).

Effect of Organic Solvents on the Stability of Protease To investigate the stability of strain K protease in organic solvents, the cell free supernatant of the culture was incubated at 37°C and agitated at 150 rpm with 25% (v/v) of organic solvent for 14 days. When 1-pentanol, benzene, toluene, p-xylene and n-hexane, of which the log Po/w values were between 1.3 to 3.5, was added to the supernatant of the culture, the proteolytic enzyme of strain K was inactivated more than 35% compared to the control. When organic solvents such as 1-decanol, isooctane, decane, dodecane and hexadecane with log Po/w values between 4.0 and 8.8 were added to the cell free supernatant, the enzyme activities were respectively 1.10, 1.59, 1.20, 1.49 and 1.52 times more stable compared to the control. As can be seen in the profile (Figure 4), the protease produced by Pseudomonas aeruginosa strain K was activated in the presence of organic solvents, especially with log Po/w values equal or more than 4.0. Figure 4 shows the relative activity of protease in the presence of organic solvent compared to the non-solvent containing control. The effect of different percentages of various organic solvents for Pseudomonas aeruginosa strain K was studied. As shown in the stability profile (Figure 4), strain K protease was 2.59, 1.59 and 1.20 times more active in the presence of 75% (v/v) of 1-decanol, isooctane and ndodecane, respectively. Whereas, this proteolytic enzyme was partially inactivated by other organic solvents tested at 75% (v/v). It was observed that strain K protease was stable in 50% (v/v) of benzene, n-hexane, 1-decanol, isooctane and n-hexadecane (Figure 4). Likewise, the

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enzyme was found to be stable in 25% (v/v) of 1-decanol, isooctane, n-decane, n-dodecane and n-hexadecane. Therefore, the stability data could be interpreted as differential solventtolerant of different isomers of strain K protease (Geok et al., 2003).

3.5

Relative stability

3 2.5 2 1.5 1 0.5

1-

Pe nt an

ol Be nz en e To lu en e pXy le ne nHe xa ne 1De ca no l Is oo ct an e nDe ca ne nDo de ca ne H ex ad ec an e

0

Organic solvents (% v/v)

Figure 4: Effect of Different Percentages of Organic Solvents on the Stability of Strain K Protease. The cell-free supernatant was incubated at 37°C with shaking 150 rpm for 30 min in the presence of (v/v) 0% ( ), 25% ( ), 50% ( ) and 75% ( ) organic solvents (Source: Geok et al., 2003).

Relative activity (%)

250 200 150 100 50

C

on t

ro l

(c el

l- f re e

su pe

rn a Et hy t an t) la ce ta Be t e nz en 1e H ep ta no l To lu Et e ne hy lb en ze ne H ex an 1e D ec an ol Is oo ct nan D e od e nc Te a tr a ne de ca ne

0

Solvents (25% v/v) Figure 5: Effect of various organic solvents on the stability of crude protease activity from B. cereus strain 146.

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Effects of various organic solvents on the stability of crude protease activity of Bacillus cereus strain 146 was investigated (Shafee et al., 2006). Activity of the protease was drastically increased in the presence of 1-decanol, isooctane, n-dodecane and n-tetradecane, but reduced in the presence of ethyl acetate, benzene, toluene, 1-heptanol, ethylbenzene and hexane (Figure 5).

Purification of Organic Solvent-tolerant Protease Pseudomonas aeruginosa strain K extracellular protease was purified to homogeneity by ammonium sulphate precipitation and anion exchange chromatography (Rahman et al., 2006). Results of the purification of P. aeruginosa strain K extracellular protease were summarized in Table 3. Three protease peaks were eluted from DEAE Sephacel column, suggesting that this bacterium produced three types of proteases. Based on the stability test results (Figure 6), the enzyme represented by peak 2 (P2) was inactivated or denatured in the presence of organic solvents. However, the enzymes represented by peak 1 (P1) and 3 (P3) were proven to be tolerant to 25% (v/v) isooctane, decane and dodecane. P3 enzyme was apparently more organic solvent-tolerant than P1 enzyme. Therefore, the ability of this proteolytic enzyme to tolerate high percentages of organic solvents was chosen for characterization studies. The organic solvent-tolerant protease (P3) was purified to homogeneity with a total yield of 39.91% and 124.43 folds (Table 3). The organic solvent-tolerant protease K from Peak 1 was also purified using a different method. The protease was purified to 96.1 folds with a yield of 4.6% using ultrafiltration (5000MW cutoff) and hydrophobic interaction chromatography techniques. Table 3: Purification Table of Pseudomonas aeruginosa strain K Protease Purification Step Crude (NH4)2SO4 Ppt

Total Activity (U) 7424.00 3529.28

DEAE Sephacel Chromatography Fraction 1 (P1) 1868.62 Fraction 2 (P2) 1418.73 Fraction 3 (P3) 2962.92

Total Protein (mg) 56.12 1.41

Specific Activity (U/mg) 132.29 2503.04

Fold

Recovery (%)

1.00 18.92

100.00 47.54

0.62 0.53 0.18

3013.90 2676.85 16460.67

22.78 20.23 124.43

25.17 19.11 39.91

Note: Protease activity was assayed by the method of Keay and Wildi (1970). One unit (U) of protease activity is equivalent to 0.5 µg tyrosine liberated by 1.0 ml of enzyme solution under the assay conditions. Protein concentration was determined by measuring absorbance at 280 nm using bovine serum albumin as a standard (Source: Rahman et al., 2006).

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2

Relative activity (%)

1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0 P1

P2

P3

Peak Figure 6: Organic solvent stability of enzyme from Peak 1, Peak 2 and Peak 3, which represented by P1, P2 and P3, respectively in 25% (v/v) isooctane ( ), decane ( ) and dodecane ( ). Relative activity was calculated with reference to the activity of the enzyme incubated in the absence of organic solvent.

Characterization of Organic Solvent-tolerant Protease The molecular weight of the purified organic solvent-tolerant protease K from peak 3 was determined to be about 51kDa by SDS-PAGE, while the enzyme from peak 1 was determined to be 30 kDa. Strain K protease (peak 3) was found to be active on a broad range of pH values between 5.0 to 11.0 at 37°C with the maximum pH of 10.0 for hydrolysis of casein (Table 4). The activity was reduced approximately 40% if the pH was maintained at pH 11.0. Based on this observation, strain K protease could be classified as an alkaline protease. The protease activity of the purified enzyme (peak 3) was measured at temperatures ranging from 37°C to 80°C. Optimum activity of the purified enzyme was exhibited at 70°C (Table 4) while about 3% of the activity was inactivated after exposure to 80°C for 10 min. The optimum temperature studies indicated that strain K protease was active over a fairly wide range of temperature values from 37 to 80°C. The thermostability study of strain K protease was carried out at temperature ranging from 37°C to 80°C. The purified enzyme appeared to be stable and retained its full activity after 30 min incubation from 37°C to 50°C. Proteolytic activity decreased dramatically when the temperature increased above 70°C with only 38% and 2% of activity remaining at 70°C and 80°C, respectively. Therefore, it could be hypothesized that higher optimum temperature does not necessarily lead to higher thermostability. The half-life of strain K protease at 50°C was estimated to be 36 h. The thermostability profile at 37°C showed that 60% and 40% of its protease activity still remained after 96 h and 120 h, respectively. Therefore, the estimated half-life of strain K protease at 37°C was 108 h.

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Enzyme inhibition experiments showed that phenylmethylsulfonyl fluoride (PMSF), which is an inhibitor of serine proteases, was the most effective agent in the activating the strain K protease (Table 4). The protease strain K was identified as a metalloprotease due to its inhibition towards metal chelators such as EDTA and o-phenantroline. The protease was activated by Zn2+ and Sr2+ while Fe3+ inhibited it. Refering to Table 4, activation effect was also observed in denaturing and reducing agents such as 6M urea, Triton-C-100 and Tween 20. Strain K protease was capable of hydrolyzing all the soluble and insoluble substrates tested except bovine albumin. In this study (Table 4), the purified casein showed the highest degree of susceptibility to proteolysis. Whilst, the insoluble protein substrate (hemoglobin) was less susceptible compared to both the purified and hammansten caseins. This protease also showed the ability to hydrolyze large molecules such as azocoll. Table 4: Summary of characterization of the organic solvent-tolerant strain K protease

Tested Parameter Temperature pH Inhibitors Metal I ons Additives Substrates

Activating agent 70°C 10.0 Phenylmethylsulfonyl fluoride (PMSF) Zn2+, Sr2+ Urea, Triton-C-100, Tween 20 Purified casein, hammansten casein, azocoll, hemoglobin

The enzyme was stable to all the organic solvents tested after 30 min incubation. Organic solvents such as isooctane, n-hexadecane, n-dodecane and n-decane (25% v/v) enhanced the stability of purified strain K protease by 85%, 83%, 78% and 59%, respectively when compared to the control (Figure 7). As shown in the profiles (Figure 7), the purified protease produced by Pseudomonas aeruginosa strain K was activated in the presence of organic solvents, especially with log Po/w values equal or more than 4.0 (1-decanol, isooctane, ndecane, n-dodecane and n-hexadecane).

Analysis of the Nucleotide Sequence of the Organic Solvent-tolerant Protease Gene The gene of organic solvent-tolerant protease from Pseudomonas aeruginosa strain K was amplified by polymerase chain reaction using consensus primers based on the multiple sequence alignment of alkaline and metalloprotease genes from Pseudomonas species. Nucleotide sequence analysis of the gene revealed an open reading frame containing 1440bp, which encodes for a polypeptide of 479 amino acid residues. The polypeptide composed of an N-terminal propeptide of 7 amino acid residues and a mature protein of 472 amino acid residues. The gene shared high homology with alkaline and metalloprotease sequences from

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P. aeruginosa and P. fluorescens. The recombinant strain K protease was successfully expressed in pGEX-4T-1 expression vector. In the presence of 1mM IPTG, the recombinant strain K protease was released into the periplasm of the Escherichia coli BL21(DE3) host (Geok, 2002).

ta nt 1) Pe nt an ol Be nz en e To lu en e pXy le ne nHe xa ne 1De ca no l Is oo ct an e nDe ca nne Do de nc an He e xa de ca ne

N

on e

(c el l-f re e

su p

er na

Relative stability

2 1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0

Organic solvents (25% v/v)

Figure 7: Organic solvent stability of the purified strain K protease in organic solvents. One ml of organic solvent was added to 3.0 ml of the purified protease and incubated at 37°C, with a shaking rate of 150 rpm for 14 days. The remaining proteolytic activity relative to the solvent-free control (0% v/v) was shown as the stability.

The complete nucleotide sequence of the organic solvent protease 115b showed an open reading frame (ORF) of 1149 that encoded a polypeptide of 383 amino acid residues and the calculated molecular mass of the protein was 39,448 Da (Mahamad, 2006). ORF also encoded a single peptide consisting of 29 residues and a propeptide of 79 residues. The mature protein comprised 275 amino acids with a calculated molecular mass of 27,846 Da. The amino acid residues from Bacillus pumilus 115b protease showed high homology (90%) with alkaline serine protease from B. pumilus TYO-67 and B. pumilus UN-31-C-42. To date, the organic solvent tolerant protease K (wild-type) which exhibited a tremendous stability in many of the organic solvents tested was fully purified and well characterized. On the other hand, the purification and characterization of two (B. pumilus 115b protease and B. cereus strain 146) other organic solvent tolerant proteases are in progress.

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REFERENCE Allen, K. N., Bellamacina, C. R., Ding, X., Jeffery, C. J., Mattos, C., Petsko, G. A., and Ringe, D., 1996. An experimental approach to mapping the binding surfaces of crystalline proteins. Journal of Phyical Chemistry, 100: 2605–2611. Cruden, D. L., Wolfrom, J. H., Rogers, R. D., and Gibson, D. T., 1992. Physiological properties of a Pseudomonas strain which grows with p-xylene in a two-phase (organicaqueous) medium. Applied and Environmental Microbiology, 58: 2723-2729. Fitzpatrick, P. A., Steinmetz, A. C. U., Ringe, D., and Klibanov, A. M., 1993. Enzyme crystal structure in neat organic solvent. Proceedings of The National Academy of Sciences of USA, 90: 8653-8657. Geok, L. P., 2003. Production, purification and characterization and expression of organic solvent tolerant protease gene. PhD Thesis. Universiti Putra Malaysia. Geok, L. P., Razak, C. N. A., Rahman, R. N. Z. A., Basri, M., and Salleh, A. B., 2003. Isolation and screening of an extracellular organic solvent-tolerant protease producer. Biochemical Engineering Journal, 13: 73-77. Ghorbel, B., Sellami-Kamoun, A., and Nasri, M., 2003. Stability studies of protease from Bacillus cereus BG1. Enzyme and Microbiol Technology, 32: 513–518. Gupta, A., Roy, I., Khare, S. K., and Gupta, M. N., 2005. Purification and characterization of a solvent stable protease from Pseudomonas aeruginosa PseA. Journal of Chromatography, 1069: 1069: 155–161. Keay, L., and Wildi, B. S., 1970. Proteases of Genus Bacillus. I: Neutral Proteases. Biotechnology and Bioengineering, 12: 179-212. Klibanov, A. M., 2001. Improving enzymes by using them in organic solvents. Nature, 409: 241-246. Mahamad, S., 2006. Cloning, sequencing and expression of an organic solvent tolerant protease from Bacillus pumillus 115b. M. S Thesis. Universiti Putra Malaysia. Ogino, H., Yasui, K., Shiotani, T., Ishihara, T., and Ishikawa, H., 1995. Organic solventtolerant bacterium which secretes an organic solvent-stable proteolytic enzyme. Applied and Environmental Microbiology, 61: 4258–4262. Ogino, H., Watanabe, F., Yamada, M., Nakagawa, S., Hirose, T., Noguchi, A., Yasuda, M., and Ishikawa, H., 1999. Purification and characterization of organic solvent-stable protease from organic solvent-tolerant Pseudomonas aeruginosa PST-01. Journal of Bioscience and Bioengineering, 87: 61-68. Ogino, H., Uchiho, T., Yokoo, J., Kobayashi, R., Ichise, R., and Ishikawa, H., 2001. Role of intermolecular disulfide bonds of the organic solvent-stable PST-01 protease in its organic solvent stability. Applied and Environmental Microbiology, 67: 942–947. Rahman, R. N. Z. A., Geok, L. P., Basri, M., and Salleh, A. B., 2005a. Physical factors affecting the production of organic solvent-tolerant protease by Pseudomonas aeruginosa strain K. Bioresource Technology, 96: 429–436. Rahman, R. N. Z. A., Geok, L. P., Basri, M., and Salleh, A. B., 2005b. An organic solventtolerant protease from Pseudomonas aeruginosa strain K Nutritional factors affecting protease production. Enzyme and Microbiol Technology, 36: 749-757.

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Rahman, R. N. Z. A., Geok, L. P., Basri, M., and Salleh, A. B., 2006. An organic solventstable alkaline protease from Pseudomonas aeruginosa strain K: enzyme purification and characterization. Enzyme and Microbial Technology, (In press). Schmitke, J. L., Stern, L. J., and Klibanov, A. M., 1998. Comparison of x-ray crystal structures of an acyl-enzyme intermediate of subtilisin Carlsberg formed in anhydrous acetonitrile and in water. Proceedings of The National Academy of Sciences USA, 95: 12918-12923. Shafee, N., Aris, S. N., Rahman, R. N. Z. A., Basri, M., and Salleh, A. B., 2005. Optimization of environmental and nutritional conditions for the production of alkaline protease by a newly isolated bacterium Bacillus cereus strain 146. Journal of Applied Sciences Research, 1: 1-8. Shafee, N., Tan, C-C., Mahamad, S., Rahman, R. N. Z. A., Basri, M., and Salleh, A. B., 2006. Nutritional factors affecting organic solvent-tolerant alkaline protease production by a new Bacillus cereus strain 146. Annals of Microbiology, 56: 29-34. Singh, J., Batra, N., and Sobti, R. C., 2000. Serine alkaline protease from a newly isolated Bacillus sp. SSR 1. Process Biochemistry, 36: 781-785. Tanaka, A., and Kawamoto, T., 1991. Immobilized enzymes in organic solvents. Bioprocess Technology, 14: 183-208. Yennawar, N. H., Yennawar, H. P., and Farber, G. K., 1994. X-ray crystal structure of gamma-chymotrypsin in hexane. Biochemistry, 33: 7326-7336.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 111-125

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 7

IMMOBILIZED ENZYMES Mohd Basyaruddin Abdul Rahman, Noor Mona Md. Yunus, Siti Salhah Othman, Abu Bakar Salleh and Mahiran Basri ABSTRACT The use of immobilized enzymes as biocatalysts in organic syntheses is of increasing interest. Immobilized enzymes are enzymes attached to carriers via physical or chemical interactions, thus making them to be easily handled as compared to their soluble counterparts. The first part of the chapter outlines the advantages immobilization, different immobilization techniques, types of supports and the uses of immobilized enzymes. The latter part of the chapter reviews our work in immobilizing lipase onto various supports. Types of supports that have been studied are hydrophobic, hydrophilic, advanced material and natural supports. The immobilized lipases prepared onto different supports were characterized in term of their activity, thermostability and enantioselectivity. Hydrophobic lipases exhibited relatively higher activities when immobilized on the more polar polymers (Amberlite XAD7, XAD8 and RCOOH). The activities of immobilized lipase onto hydrogels were increased when the hydrophilicity was increased. Immobilization onto synthetic clay-like material of hydrotalcite (Mg/Al) gave high activity and enantioselectivity. The use of the hydrotalcite intercalated with Sodium Dodecyl Sulfate (SDS) showed an increase in adsorption of lipase as well as the lipase activity. Immobilized lipase onto inexpensive natural kaolin exhibited increased activity by four fold when compared to native lipase.

INTRODUCTION Immobilization of enzymes is a technique to localize enzymes, the catalytic functional component which converts the substrate of interest into the desired products onto a carrier, the non-catalytic structural component to aid separation and reuse of the catalyst (Cao et al., 2003).

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Recently, increasing emphasis on the use of biocatalysts in synthetic reactions has resulted in a similar increase in the use of immobilized enzymes, thus making this technique a basis for numerous biotechnological processes as well as analytical devices.

AIM AND ADVANTAGES OF IMMOBILIZATION Since the beginning of seventies, immobilized enzymes has been one of the exciting products of biotechnology as its offers quite a number of advantages over their soluble counterparts in terms of stability, reusability, applicability to continuous processing, minimization of pH and substrate-inhibition effects (Pal et al., 2000). Immobilized enzyme may be specific for certain types of substrate thus would lead to the formation of specific compounds, high yields, possibility of working under mild operating conditions and less contamination for both the final product and the environment (Dumitriu et al., 2003; Garcia et al., 2000; Jonzo et al., 2000). Immobilized lipases are particularly interesting for industrial purposes since they are easily handled. Unlike soluble lipases, lipases in immobilized form are strongly bound to a solid phase, making it easy to separate the catalyst from the reaction mixture. Besides, immobilized enzyme makes continuous process possible, improves operational performance of the enzyme and allows enzyme recycling, favouring continuous reactor operation (Jeison et al., 2003; Salis et al., 2003; Garcia et al., 2002). Another aim of immobilizing enzymes is to encourage enzymes functional activities as well as to stabilize enzymes and proteins. Stabilization of enzymes can be achieved when the chemical and/or physical properties of the enzymes is altered as desired through physical interactions between the supports and the enzymes. Through immobilization, enzymes are dispersed over a larger surface area, which allows exposure of enzymes to the more homogeneous substrate concentration. The extent of stabilization depends on the enzyme structure, the immobilization method and type of supports (Bayramoglu et al., 2005). Immobilization of enzymes may protect the enzyme to some extent from solvent denaturation (Dalla-Vecchia et al., 2005). For example, the immobilization of brain glutamine synthetase on a liquid chromatographic stationary phase increased the stability of the enzyme from hours to weeks (Bartolini et al., 2003; Cloix and Wainer, 2001). Now immobilization becomes much more than a need, it also becomes a powerful tool to improve enzyme selectivity. In general, we can assume that any biotransformation where the enzyme is undergoing important conformational changes during catalysis should be strongly modulated via the design of controlled protocols of enzyme immobilization. The immobilization of the enzymes through areas more or less connected or related to the active center should promote attentions on those conformational changes and hence, in the activity and selectivity of the enzyme ( Fernandez-Lorento et al., 2001). Immobilization methods and matrixes used could have profound effects on the resulting biological activity of the bound enzymes. Meanwhile, the immobilization method could also affect the enzyme activity through the chemical modification of the amino acids involved in coupling steps especially when the coupling site was much closer to the active site of the

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enzyme. A suitable matrix and immobilization method are therefore the key points in enzyme immobilization (Xi et al., 2005; Bayramoglu et al., 2004).

TYPES OF IMMOBILIZATION Enzyme immobilization can be accomplished in different ways, and these may be divided into two main categories, physical methods and chemical methods. Physical methods such as adsorption based on weaker interaction between the enzyme and solid support while chemical methods based on formation of covalent attachment to a carrier (Chiou and Wu, 2004; Arica et al., 2001; Persson et al., 2000). Recently, many immobilization procedures have been developed as shown in Figure 1. Among these are physical adsorption, covalent binding, encapsulation, entrapment within a matrix of a polymer and cross linking (Noureddini et al., 2005; Salis et al., 2003).

Physical Adsorption

Covalent Binding

Encapsulation

Entrapment

Support Enzyme molecule Membrane Cross linking

Cross linking polymer

Figure 1: Immobilization of enzyme

The main tasks when designing immobilized biocatalyst are to select suitable carrier, immobilization conditions (pH, temperature and nature of medium) and the enzyme itself (source, nature and purity). Many supports, such as polymers and resins, silica and silicaalumina composites and carbonaceous materials, mostly in the form of powders, beads and chips have been studied (Cao, 2005). These systems are often severely diffusion limited, leading to considerable fraction of unused enzymatic activity. A new monolith catalyst

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support with a very open wall structure was used for enzyme-catalyzed reactions (Lathouder et al., 2004). In recent years, there has been an increasing trend of using immobilization techniques as a complementary tool to protein engineering. By exploring or combining available methods or knowledge or by exploiting the positive attributes to each immobilization methods, the performance of immobilized enzymes can be improved to levels that were previously unattainable by use of single immobilization methods (Cao, 2005). For example, Ma et al., (2004) had describe a method whereby a lipase enzyme porcine pancreatic lipase, PPL, was first physically adsorbed in the channels of an MCM-41 support, and the size of the channels’ mouth was subsequently reduced by chemical modification by covalent coupling with an organic siloxane. This produced pores with narrow necks and wide bodies which prevented leaching of the weakly bound immobilized enzyme, but did not inhibit access of substrate to the enzyme.

Physical Adsorption Adsorption may have a higher commercial potential than other methods because adsorption is simpler and less expensive, and a high catalytic activity may be retained. The method also offers the reusability of expensive supports after inactivation of immobilized enzymes. However, adsorption generally is not very strong, and some of the adsorbed protein will be desorbed during washing and operation. Thus, enzyme immobilization via adsorption requires a strong hydrophobic or ionic interaction between the enzyme and support (Bayramoglu et al., 2002). The hydrophobic group density, pH and temperature can have important effects on the adsorption of lipase on the support. The properties of the support seem to provide an adequate approach for the adsorption of lipase based on their hydrophobic amino acid content. The phenylalanine containing poly (HEMA-MAPA) membrane revealed good properties as an adsorptive membrane and is useful for enzyme immobilization via hydrophobic interaction (Arica et al., 2001). The enzyme could be repeatedly adsorbed and desorbed without any significant loss in adsorption capacity. In addition, the reusability of the membrane support may provide economic advantages for large scale biotechnological approaches. Physical adsorption is an appropriate immobilization method because it promotes a strong but reversible immobilization that enables support reusability after easy removal of the spent enzyme (Salis et al., 2003). Physical adsorption results in minimal disruption of the structure of the immobilized enzyme but loss of the enzyme immobilized on the support is unavoidable due to the weak bonds between the enzyme and the support (Ma et al., 2004).

Encapsulation and Entrapment Other techniques of immobilization are the encapsulation and entrapment methods. The immobilization is achieved by inclusion of the enzyme molecule within a gel matrix of a cross-linked polymer. Immobilization is done by mixing the enzyme with the monomer or soluble polymer followed by cross-linking to form a gel. The result of this is that the enzyme

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can move freely in the solution but movement is restricted within the interstitial spaces of the gel. Commonly used matrix for enzyme encapsulation is sol-gel and in some cases, it was even found to increase the stability of the enzymes and proteins. Substrates diffuse into the network of pores and channels of the sol-gel matrix to reach the active sites of the encapsulated enzymes to produce the product. However, the disordered network pores and channels in the sol-gel matrix limits the reactions allowed. For this reason, Han et al., (2002) have expanded the enzyme encapsulation technique by utilizing the advantage of the large pore size structure of the mesoporous silicate material, mesocellular form, MCF to encapsulated FeHeme chloroperoxidase (CPO) from Caldariomyces fumago. Several chemical approaches have been exploited to incorporate enzyme molecules either on fiber surfaces or inside the fibers. These fibrous materials offer versatile porosity and ultrahigh specific surface for superior enzyme binding and substrate accessibility. Several synthesis and processing concepts have been explored to create specific surface properties targeted to bind enzyme proteins.

Crosslinking Method In the crosslinking process, glutaraldehyde is the most common reagents used to crosslink the enzyme molecules. Bartolini et al., 2003 reported that the inclusion of glutaraldehyde to immobilize glycolytic enzyme glyceraldehydes-3-phosphate dehydrogenase (GAPDH) to silica matrix showed an increase in the stability of the GAPDH. Furthermore, packed GAPDH-Glut-P in a chromatographic column to make immobilized enzyme reactor (IMER) preserved the enzymatic activity from inactivating processes, widely increasing the stability of the enzyme with advantages in term of accuracy and reproducibility. As such, the immobilized enzyme can be placed in an on-line system for rapid screening of compounds for inhibitory activity. Bayramoglu et al., (2005) reported that immobilization of lipase onto 1,6-diaminohexane (as a spacer arm) of poly (GMA-HEMA-EGDMA) using glutaric dialdehyde as the coupling reagent resulted in an increase in the apparent activity of lipase covalently attached to the epoxy group of the support. The immobilized lipase onto the spacer arm retained much activity over wider ranges of temperature and pH, indicating that immobilized enzymes become less sensitive to reaction condition. Moreover, the high operational stability showed that immobilized enzyme can be used in the continuous system. Another coupling agent used in immobilization process was 1-Ethyl-3-(3-dimethylaminoprpyl) carbodiimide hydrochloride, C8H17N3.HCl (EDC), to attach lipase to chitosan supports containing hydroxyl groups which was done by Chiou and Wu, (2004). They found immobilization enhanced the enzyme stability against changes of pH and temperature. High storage stability of 30 days and an increase enzyme activity were observed and the activity retained its initial activity after 10 batch hydrolytic cycles. Recently, Ragheb et al., (2005) reported their work on a simple lipase entrapment in silicone rubber using tetraethyl orthosilicate as a crosslinker, to give a highly active silicone-enzyme elastomer.

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Covalent Binding Covalent attachment is an important immobilization method for enzymes which has been demonstrated to increase higher resistance to heat, denaturants, pH extreme and organic solvent (Fernades et al., 2004). Martin et al., (2003) reported that covalent attachment of Thermoanaerabacter sp. CGtase onto Eupergit C exhibited an improved resistance against thermal (half life of the immobilized enzyme at 95oC was five times higher than that of the soluble enzyme) and pH denaturation, and has promising operational stability for the production of maltooligosaccharides. Epoxy supports are almost-ideal for very easy covalent immobilization of enzymes on both the laboratory and industrial scale. The epoxy supports are also able to form very stable covalent linkages with different protein groups (amino, thiol, phenolic types) under very mild experimental condition. For example, Bayramoglu et al., (2004) reported their work on immobilized α-amylase onto polyhydroxyethylmethacrylate-glycidyl methacrylate via amide linkage between amino group of α-amylase and the epoxy groups of the supports. The immobilized α-amylase showed higher thermostability with 4% initial activity loss during continuous operation for 120 h. Huang et al., (2003) has opened the opportunity of immobilizing enzyme onto magnetic nanoparticles. Lipase was covalently bound onto Fe3O4 magnetic nanoparticles (12.7 nm) via carbodiimide activation. Magnetic measurement revealed the resultant lipase-bound magnetic nanoparticles were superparamagnetic. Compared to the free enzyme, the bound lipase exhibited a 1.41-fold enhanced activity, a 31-fold improved stability, and better tolerance to the variation of solution pH. It revealed that the available active sites of lipase and their affinity to substrate increased after being bound onto magnetic nanoparticles. The first covalent immobilization of CPO chloroperoxidase isolated from Caldariomyces fumago was onto epoxide derivatized silica gel. The immobilization was performed under mild conditions. The enzyme was still active after the immobilization process; no leaching of the enzyme from the support was detectable after repeated washings. The immobilization of CPO enhanced the stability of the enzyme with respect to the effect of pH and oxidizing agent concentration (Petri et al., 2004).

NATURE OF SUPPORTS The search for the most effective procedures for immobilizing enzymes has led to the use of different kinds of supports. The ideal supports would increase substrate binding, decrease product inhibition, shift the apparent pH optimum to the desired value, increase enzyme stability, discourage bacterial growth and enable the enzyme to be readily recovered for reuse. At the same time, cost of production and ease of preparation must also be considered. The supports used for the immobilization must provide specific morphological and chemical features, like particle size, pore size and specific surface area. The chemical nature of the support surface can influence water partitioning, substrate/product inhibition and enzyme activity (Salis et al., 2003). The physical structure and chemical composition of support can also influence the microenvironment of the immobilized enzymes and consequently their biological properties (Bayramoglu et al., 2004).

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The matrices used for enzyme coupling may be roughly divided into two groups, namely artificial and natural polymers. Generally, artificial polymers such as polyacrylamide, polyacrylate and polystyrene are resistant to pressure but they had disadvantage of low compatibility and might cause enzyme denaturing resulting from hydrophobic interaction between the polymer and the enzyme. Natural polymers such as agarose, cellulose and crosslinked dextran have high compatibility and low protein unfolding effect, but they cannot resist high operation pressure and the deep inner macroporous network of the beads would trap the enzyme, which might leach from matrix under operational conditions (Xi et al., 2005). The morphology of the support such as surface hydrophobicity, charge and chemical modification of the surface material can affect both the stability and the orientation of a protein at a surface and may significantly affect the reaction rate and/or selectivity of the process (Xi et al., 2005; Plou et al., 2002). Mesoporous silicates with large-pore size structure, such as SBA and mesocellular foam, MCF load more enzymes and the large porosity of the materials provide better access for the substrates to the immobilized enzymes (Han et al., 2002). Generally, the efficiency of enzyme immobilization is decided by the support size (or surface area). However, this is questionable in the case of lipase, because of the explicit structure of the enzyme protein (Chiou and Wu, 2004). Activation of a mesoporous silica having high surface area with octyltriethoxysilane has proven successful to cover the hydrophilic surface of silica with a hydrophobic layer of octyl groups. This has enabled the objective to achieve the immobilization of lipase in a monolayer, avoiding the formation of enzyme aggregates. A disordered network of pores and channels, may only allow the smallest substrate to penetrate while bigger substrates would clog the channels, may only allow the smallest substrates to penetrate while bigger substrates would clog the channels, slowing the reactions. Thus a heterogeneous pore size distribution interferes not only with the optimization of enzyme load on the internal surfaces, but also with the diffusion of substrates and/or products. In general, the wider the pore size is, the lower the surface area and hence the enzyme loading should be lower for an equivalent density of active groups available for the enzyme per area unit. Thus, catalytic efficiency depends on the equilibrium of monolayer enzyme arrangement, pore size assuming homogeneous distribution and surface area (Blanco et al., 2004).

OUR WORK Immobilization of lipase on various supports was carried out in our laboratory. Most of the work done is focused on the use of lipase from Candida rugosa, a commercially available lipase (Sánchez et al., 1999) with interesting properties such as high stereo-, enantio-, regioand substrate specificity (Balcao et al., 1996).

Hydrophobic Support The lipase was immobilized onto various supports using various methods to enhance their catalytic properties. Amongst the first attempts to produce efficient and more stable lipase was by attaching various hydrophobic groups (monomethoxy polyethylene glycol,

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acetyldehyde, dodecyldehyde, methyl acetonimidate and methyl 4-phenylbutyrimidate) on the enzyme molecules through chemical modification (Ampon et al., 1994). This was according to the general principle whereby; strong hydrophobic or electrostatic interaction between enzyme and support is important for a successful adsorption in an immobilization procedure (Basri et al., 1994a). These derivatised lipases were then immobilized on various supports. The strong hydrophobic interactions can be achieved by using hydrophobic supports or derivatised lipases. Lipase modified using 45 % of monomethoxy polyethylene glycol with molecular weight of 1900 was chosen for further immobilization on polymer beads of different polarity such as Amberlite XAD2 and XAD4 (non-polar), Amberlite XAD7 and XAD8 (medium polar) and polar polycarboxylic acid and polyacrylonitrile. Figure 2 shows the SEM image of immobilized lipase onto Amberlite XAD7.

Amberlite XAD7-lipase

Figure 2: SEM image of immobilized lipase onto Amberlite XAD7.

As expected, hydrophobicity of lipase plays an important role towards increasing the amount of immobilized activity due to the stronger preferential binding of the more hydrophobic lipase on the supports. The hydrophobic lipases also exhibited relatively higher activities when immobilized on more polar polymers (Amberlite XAD7, XAD8 and RCOOH). This can be explained by the fact that the polarity of these beads facilitates ionic interaction, in addition to the existing hydrophobic interaction of the enzymes which further strengthened the overall interaction. In the study, it was also found that the activity of lipase adsorbed on smaller beads was higher compared to larger ones in the esterification of propyl oleate. This can be explained by the fact that smaller beads adsorbed considerably more enzymes before the saturation point of adsorption was reached due to the greater surface area (Basri et al., 1994b).

Hydrophilic Support Immobilization of lipase through entrapment on poly(N-vinyl-2-pyrrolidone-co-2hydroxyethyl methacrylate) hydrogel had also been extensively studied by Basri and coworkers (1999). In this study, hydrogels of varying composition of monomers, N-vinyl-2-

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pyrrolidone and 2-hydroxyethylmethacrylate, did not affect the activity of lipase but the presence of unreacted monomers decreased the activity of the lipase to less than 50 % in the esterification of oleic acid and butanol in hexane. Apparently, the reactive/unreacted monomers may have a poisoning effect on the enzyme, thus decreasing its activity. Increasing the percentage of N-vinyl-2-pyrrolidone from 0 to 90, which corresponded to an increase in hydrophilicity of the hydrogel and its equilibrium water content, seemed to increase the activity of the immobilized enzyme. Hydrogel with 90: 10 composition of Nvinyl-2-pyrrolidone: 2-hydroxyethylmethacrylate, was the best support for immobilization as it gave the highest activity owing to the sufficient water surrounding the enzyme in the hydrogel. In contrast, low activity was observed for lipase immobilized on 100 % 2hydroxyethylmethacrylate. This may be due to the decrease in equilibrium water content of 2hydroxyethylmethacrylate or partial solubility of this polymer in water and lower level of crosslinking resulting in diffusion of lipase from the gel.

Advanced Material Support The search for suitable immobilization supports was continued by immobilizing lipase on a synthetic clay-like material, called hydrotalcite (Othman et al., 2003). Hydrotalcite is classified under the layered double hydroxides group of minerals, with anion exchange capacity and extensive areas of reactive hydroxylated surfaces (Miyata and Hirose, 1978). It exists in various types ranging from Mg/Al, Ca/Al, Zn/Al, Zn/Cr and Co/Al (Oriakhi et al., 1996), which carries the mineralogical and technical importance. Interestingly, engineering and expansion of its pore size is possible by varying the composition of cations, by heat treatment or through intercalation of surfactants into the lamellar spaces. Amongst the various types, the Mg/Al type of hydrotalcite is the most common member of the layered double hydroxides minerals (Carlino and Hudson, 1995). In our work Mg/Alhydrotalcite was prepared at various ratios of Mg: Al and comparison of their lipase adsorption capacity and enhancement of lipase activity in the esterification of butyl oleate was carried out. Mg/Al-hydrotalcite prepared at ratio 4 exhibited better lipase loading and catalytic activity. Abdul Rahman et al., (2004a) showed that the application of layered double hydroxide of Mg/Al and its nanocomposites Mg/Al-sodium dodecyl sulphate (SDS) as support for enzyme immobilization have exhibited an increase in the stability and activity of the enzyme. The XRD spectra showed that basal spacing for synthesized Mg/Al and Mg/Al-SDS were around 7.8 and 34.3 Å, respectively. The expansion of layered structure was observed to accommodate the surfactant anion between the interlayer for the Mg/Al-SDS. The increase of the BET surface area of the support from 60 to 71 m2/g has increased the amount of protein adsorbed on Mg/Al and Mg/Al-SDS from 24969 and 47861 mg, respectively. Higher amount of protein adsorbed (70.8%) on Mg/Al-SDS was due to the larger surface area and thus increasing the specific surface area of the Mg/Al-SDS support as compared to protein adsorbed (36.9%) on Mg/Al. Similar findings also have been observed by Abdul Rahman et al., (2004b) when using Zn/Al and Zn/Al-dodecyl sodium sulfoacetate (DSS) nanocomposite. High percentage of protein adsorbed up to 78.5% in Zn/Al-DSS may be due to the large surface area of the support. During adsorption, the protein molecules were randomly scattered on the surface. Stability and enzyme activity was higher than native lipase.

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Natural Support There are varieties of commercially available supports such as polymer resin, zeolites and hydrotalcites that can be used for the immobilization of enzyme. Natural kaolin is inexpensive and shows good potential as support material. Currently, strong interests in such natural supports are due to ecofriendly demands in many modern industrial applications. Abdul Rahman et al., (2005) reported almost 77% of protein content of lipase was immobilized onto natural kaolin by physical adsorption method. In this study, lipase was immobilized onto kaolin and was physico-chemically characterized and applied in the esterification of 1butanol and oleic acid. Kaolin-lipase exhibited higher activity and stability as compared to native lipase. Table 1: Some important applications of immobilized lipases developed from our laboratory. Supports Amberlite XAD4 Amberlite XAD7

Eupergit C Eupergit C250L Mg/Al Hydrotalcites Zn/Al Hydrotalcites

Products Ethyl Caprylate Ethyl Caproate Butyl Oleate Oleyl Oleate Palm Olein Stearate Methyl Salicylate (-)-Menthyl Butyrate Dimethyl Adipate Diethyl Adipate Dibutyl Adipate Dioctyl Adipate

Applications Esters in Fragrances and Food Liquid Wax Esters in Cosmetics

Enantiomeric Esters in Pharmaceuticals Adipate Esters in Surfactants Emollients in Cosmetics

The effectiveness of an immobilization process depends on the supports used. Such increase in stability and activity performed by kaolin-immobilized lipase was favorable, and it exhibited good tolerance to conditions as required in industrial applications. Natural kaolin showed a good potential of applying natural resource as support for biocatalyst for various organic syntheses as it allows easy immobilization through a simple and inexpensive method. Currently, the immobilized lipases available from our work are given in Table 1.

CHARACTERISTICS Although a variety of supports can be obtained commercially, supports used in the immobilization technique must be able to enhance lipase characteristics such as its activity, stability and specificity. Research on immobilization of lipase with the aim of producing highly enantioselective lipase in the resolution of enantiomeric substrates was carried out (Othman, 2004). The synthesized Mg/Al-hydrotalcite together with a number of commercially available supports such as molecular sieves (10 – 20 mesh), activated carbons (50 – 150 µm granulation, 0.5 – 1.0 mm particle size) silica gel 60 (70 – 230 mesh) Eupergit

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C, Eupergit C250L and Amberlite XAD7 were tested for their efficacy as immobilization support that would not only enhance lipase stability but also lipase enantioselectivity in the resolution of (±)-menthol with butyric anhydride as the acylating agent. Table 2 shows among the immobilized lipases screened for enantioselectivity, only lipase immobilized on Mg/Al-hydrotalcite, Amberlite XAD7, Eupergit C and Eupergit C250L produced high yields of monomeric (-)-menthyl butyrate (38 – 59 %) and exhibited considerably high enantioselectivities (98 – 90 % enantiomeric excess). Supports mentioned also contributed towards high loadings of protein (28 – 80 %). Table 2: Screening of immobilized lipase for the production of (-)-menthyl butyrate. Supports Eupergit C Eupergit C250L Amberlite XAD-7 Hydrotalcite Activated C Molecular Sieves

Immobilization (%) 65.7 79.8 69.2 28.1 15.8 19.0

ee (%) 100 94.8 92.0 80.2 42.0 2.2

When the immobilized lipases were examined for the effects of various temperatures, storage and organic solvents stabilities in the enantioselective formation of (±)-menthyl butyrates, the immobilized lipases retained high catalytic activities (53 – 83 %) and showed increased stability of 2 to 3 folds compared to the native lipase. Furthermore, the immobilized lipases showed possible and efficient reuse with HT-lipase exhibiting excellent half life (t½) of 16 days. Other immobilized lipases, however, showed lower half-life, t½ = 3.5 for Amberlite XAD7-lipase and t½ = 9.0 for Eupergit C 250 L-lipase and Eupergit C-lipase. The support, Mg/Al-hydrotalcite prepared at ratio 4 of Mg:Al was further improved by heating at 200 and 400oC. Heating was expected to expand the basal spacing of the material as a result of eliminating water molecules from the interlayers of the structure. Interestingly, heating the material to 200oC resulted in expansion of its basal spacing and a two-fold increase in lipase adsorption capacity. Stability of lipase to adverse heat and in the presence of organic solvents was slightly improved when immobilized on the 200oC heat treated Mg/Al-hydrotalcite compared to the non heat treated hydrotalcite. However, further heating the material to 400oC did not seem to increase its adsorption capacity and activity nor stability of lipase adsorbed to it. This could be due to the formation of oxide ions as a result of excessive heating, which could have caused changes in the layered structure of the material or formation of a more compact arrangement. The immobilization of lipase on insoluble solid support was aimed to solve the problems encountered using soluble form of lipase. These problems include low lipase activity, low operational stability and less specificity. Results reported regarding the use of immobilized enzymes clearly exemplified the possibility of improving lipase catalytic efficacy, enantioselectivity and stability. These phenomena are related to the alteration of the conformational changes associated to the immobilization process.

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REFERENCES Abdul Rahman, M. B, Basri, M., Hussein, M. Z., Tajudin, S. M., Rahman, R. N. Z. R. A., and Salleh, A. B., 2005. Application of natural kaolin as support for the immobilization of lipase from Candida rugosa as biocatalyst for effective esterification. Applied Clay Science, 29: 111-116. Abdul Rahman, M. B, Basri, M., Hussein, M. Z., Idris, M. N. H., Rahman, R. N. Z. R. A., and Salleh, A. B., 2004a. Immobilisation of lipase from Candida rugosa on layered double hydroxides of Mg/Al and its nanocomposite as biocatalyst for the synthesis of ester. Catalysis Today, 93-95: 405-410. Abdul Rahman, M. B, Basri, M., Hussein, M. Z., Zainol, D. H., Rahman, R. N. Z. R. A., and Salleh A. B., 2004b. Immobilization of lipase from Candida rugosa on layered double hydroxides for esterification reaction. Applied Biochemistry and Biotechnology, 118: 313-32. Ampon, K., Basri, M., Wan Yunus, W. M. Z., Razak, C. N. A., and Salleh, A. B., 1994. Immobilization by adsorption of hydrophobic lipase derivatives to porous polymer beads for use in ester synthesis. Biocatalysis, 10: 341-351. Arica, M. Y., Kacar, Y., Ergene, A., and Denizli, A., 2001. Reversible immobilization of lipase on phenylalanine containing hydrogel membranes. Process Biochemistry, 36: 847854. Balcao, V. M., Paiva, A. L., and Malcata, F. X., 1996. Bioreactors with immobilized lipases: state of the art. Enzyme and Microbial Technology, 18: 392-416. Bartolini, M., Andrisono, V., and Wainer, I. W., 2003. Development and characterization of an immobilized enzyme reactor based on glyceraldehydes-3-phosphate dehydrogenase for on-line enzymatic studies. Journal of Chromatography A, 987: 331-340. Basri, M., Ampon, K., Wan Yunus, W. M. Z., Razak, C. N. A., and Salleh, A. B., 1994a. Immobilization of hydrophobic lipase derivatives onto organic polymer beads. Journal of Chemical Technology and Biotechnology, 59: 37-44. Basri, M., Ampon, K., Wan Yunus, W. M. Z., Razak, C. N. A., and Salleh, A. B., 1994b. Stability of hydrophobic lipase derivatives immobilized on to organic polymer beads. Applied Biochemistry and Biotechnology, 48: 173-183. Basri, M., Wan Yunus, W. M. Z., Yoong, W. S., Ampon, K., Razak, C. N. A., and Salleh, A. B., 1996. Immobilization of lipase from Candida rugosa on synthetic polymer beads for use in the synthesis of fatty esters. Journal of Chemical Technology and Biotechnology, 66: 69-173. Basri, M., Wong, C. C., Ahmad, M. B., Razak, C. N. A., and Salleh, A. B., 1999. Immobilization of lipase on poly(N-vinyl-2-pyrrolidone-co-2-hydroxyethyl methacrylate) hydrogel for the synthesis of butyl oleate. Journal of the American Oil Chemists Society, 76(5): 571-577. Bayramoglu, G., Kacar, Y., Denizli, A., and Arica, M. Y., 2002. Covalent immobilization of lipase onto hydrophobic group incorporated poly(2-hydroxyethyl methacrylate) based hydrophilic membrane matrix. Journal of Food Engineering, 52: 367-374. Bayramoglu, G., Kaya, B., and Arica, M. Y., 2005. Immobilization of Candida rugosa onto spacer-arm attached poly(GMA-HEMA-EGDMA) microspheres. Food Chemistry, 92: 261-268.

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Bayramoglu, G., Yilmaz, M., and Arica, M. Y., 2004. Immobilization of a thermostable αamylase onto reactive membranes: kinetics characterization and application to continuous starch hydrolysis. Food Chemistry, 84: 591-599. Blanco, R. M., Terreros, P., Fernandez-Perez, M., Otero, C., and Diaz-Gonzalez, G., 2004. Functionalization of mesoporous silica for lipase immobilization Characterization of the support and the catalysts. Journal of Molecular Catalysis B: Enzymatic, 30: 83-83. Cao, L., 2005. Immobilised enzymes: science or art? Current Opinion in Chemical Biology, 9: 217-226. Cao, L., Langen, L. V., and Sheldon, R. A., 2003. Immobilised enzymes: carrier-bound and carrier-free? Current Opinion in Biotechnology, 14: 387-394. Carlino, S., and Hudson, M. J., 1995. Thermal intercalation of layered double hydroxides: capric acid into an Mg-Al-LDH. Journal of Materials Chemistry, 5: 1433-1442. Chiou, S. H., and Wu, W. T., 2004. Immobilization of Candida rugosa lipase on chitosan with activation of the hydroxyl groups. Biomaterials, 25: 197-204. Cloix, J. F. and Wainer, I. W., 2001. Development of an immobilized brain glutamine synthetase liquid chromatographic stationary phase for on-line biochemical studies. Journal of Chromatography A, 913: 133-140. Dalla-Vecchia, R., Sebrao, D., Nascimento, M. G., and Soldi, V., 2005. Carboxymethylcellulose and poly(vinyl alcohol) used as a film support for lipases immobilization. Process Biochemistry, 40: 2677-2682. Dumitriu, E., Secundo, F., Patarin, J., and Fechete, I., 2003. Preparation and properties of lipase immobilized on MCM-36 support. Journal of Molecular Catalysis B: Enzymatic, 22: 119-133. Fernandes, K. F., Lima, C. S., Lopes, F. M., and Collins, C. H., 2004 . Properties of horseradish peroxidase immobilsed onto polyaniline. Process Biochemistry, 39: 957-962. Fernandez-Lorente, G., Fernandez-Lafuente, R., Palomo, J. M., Mateo, C. Bastida, A. Coca, J., Haramboure, T., Hernandez-Justiz, O., Terreni, M., and Guisan, J. M., 2001. Biocatalyst engineering exerts a dramatic effect on selectivity of hydrolysis catalyzed by immobilized lipases in aqueous medium. Journal of Molecular Catalysis B: Enzymatic, 11: 649-656. Garcia, R., Martinez, M., and Aracil, J., 2002. Enzymatic esterification of an acid with an epoxide using an immobilised lipase from Mucor miehei as catalyst: Optimization of the yield and isomeric excess of ester by statistical analysis. Journal of Industrial Microbiology and Biotechnology, 28: 173-179. Garcia, T., Coteron, A., Martinez, M., and Aracil, J., 2000. Kinetic model for the esterification of oleic acid and cetyl alcohol using an immobilised lipase as catalyst. Chemical Engineering Science, 55: 1411-1423. Han, Y. J., Watson, J. T., Stucky, G. D., and Butler, A., 2002. Catalytic activity of the mesoporous silicate-immobilized chloperoxidase. Journal of Molecular Catalysis B: Enzymatic, 17: 1-8. Huang, S. H, Liao, M. H., and Chen, D. H., 2003. Direct binding and characterization of lipase onto magnetic nanoparticles. Biotechnology Progress, 19(3): 1095-1100. Jeison, D., Ruiz, G., Acevedo, F., and Illanes, A. 2003. Simulation of the effect of intrinsic reaction kinetics and particle size on the behaviour of immobilized enzymes under internal diffusional restrictions and steady state operation. Process Biochemistry, 39: 393399.

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Jonzo, M. D., Hiol, A., Zagol, I., Druet, D., and Louis-Claude, C., 2000. Concentrates of DHA from fish oil by selective esterification of cholesterol by immobilized isoforms of lipase from Candida rugosa. Enzyme and Microbial Technology, 27: 443-450. Lathouder, K. M., Bakker, J., Kreutzer, M. T., Kapteijn, F., Moulijn, J. A., and Wallin, S. A., 2004. Structured reactors for enzyme immobilisation: advantages of tuning the wall morphology. Chemical Engineering Science, 59: 5027-5033. Ma, H., He, J., Evans, D. G., and Duan, X., 2004. Immobilzation of lipase in a mesoporous reactor based on MCM-41. Journal of Molecular Catalysis B: Enzymatic, 30: 209-217. Martin, M. T., Plou, F. J., Alcalde, M., and Ballesteros, A., 2003. Immobilization on Eupergit C of cyclodextrin glucosyltransferase (CGTase) and properties of the immobilized biocatalyst. Journal of Molecular Catalysis B: Enzymatic, 21:299-308. Miyata, S., and Hirose, T., 1978. Adsorption of N2, O2, CO2 and H2 on hydrotalcite-like system: Mg2+-Al3+-Fe(CN)64-. Clays Clay Minerals, 26: 441-447. Noureddini, H., Gao, X., and Philkana, R. S., 2005. Immobilized Pseudomonas cepacia lipase for biodiesel fuel production from soybean oil. Bioresource Technology, 96: 769-777. Oriakhi, C. O., Farr, I. V., and Lerner, M. M., 1996. Incorporation of poly(acrylic acid), poly(vinylsulfonate) and poly(styrenesulfonate) within layered double hydroxides. Journal of Materials Chemistry, 6: 103-107. Othman, S. S., Basri, M., Hussein, M. Z., Taufiq-Yap, Y. H., Rahman, M. B. A., Rahman, R. N. Z. R. A., and Salleh, A. B., 2003. Heat treated hydrotalcite as support for lipase immobilization. Pertanika Journal of Science and Technology. 11(2): 145-152. Othman, S. S. 2004. Immobilization of lipase from Candida rugosa onto selected matrices for use in enantioselective preparation of (-)-menthyl butyrate. PhD. Thesis. Universiti Putra Malaysia, Serdang, Malaysia. Pal, P., Datta, S., and Bhattacharya, P., 2000. Studies on the modeling and simulation of a sequential bienzymatic reaction system immobilized in emulsion liquid membrane. Biochemical Engineering Journal, 5: 89-100. Persson, M., Wehtje, E., and Adlercreutz, P., 2000. Immobilisation of lipases by adsorption and deposition: high protein loading gives lower water activity optimum. Biotechnology Letters, 22: 1571-1575. Petri, A., Gambicorti, T., and Salvadori, P., 2004. Covalent immobilization of chloroperoxidase on silica gel and properties of the immobilized biocatalyst. Journal of Molecular Catalysis B: Enzymatic, 27: 103-106. Plou, F. J., Cruces, M. A., Ferrer, M., Fuentes, G., Pastor, E., Bernabe, M., Christensen, M., Comelles, F., Parra, J. L., and Ballesteros, A., 2002. Enzymatic acylation of di- and trisaccharides with fatty acids: choosing the appropriate enzyme, support and solvent. Journal of Biotechnology, 96: 55-66. Ragheb, A. M., Hileman, O. E., and Brook, M., 2005. The use of poly(ethylene oxide) for the efficient stabilization of entrapped α-chymotrypsin in silicone elastomers: A chemometric study. Biomaterials, 26: 6973-6983. Salis, A., Sanjust, E., Solinas, V., and Monduzzi, M., 2003. Characterisation of Accurel MP 1004 polypropylene powder and its use as a support for lipase immobilisation. Journal of Molecular Catalysis B:Enzymatic. 24-25: 75-82. Sánchez, A., Ferrer, P., Serrano, A., Pernas, M. A., Valero, F., Rúa, M. L., Casas, C., and Solà, C., 1999. Characterization of the lipase and esterase multiple forms in an enzyme

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preparation from a Candida rugosa pilot-plant scale fed-batch fermentation. Enzyme and Microbial Technology, 25: 214-223. Wan Yunus, W. M. Z., Salleh, A.B., Ismail, A., Ampon, K., Razak, C. N. A., and Basri, M., 1992. Poly(Methyl Methacrylate) as a matrix for immobilization of lipase. Apllied Biochemistry and Biotechnology, 36: 97-105. Xi, F., Wu, Jianmin, Jia, Z., and Lin, X., 2005. Preparation and characterization of trypsin immobilized on silica gel supported macroporous chitosan bead. Process Biochemistry, 40: 2833-2840. Zaitsev, S. Y., Gorokhova, I. V., Kashtigo, T. V., Zintchenko, A., and Dautzenberg, H., 2003. General approach for lipases immobilization in polyelectrolyte complexes. Colloids and Surfaces A: Physicochemical Engineering Aspects, 221: 209-220.

In: New Lipases and Proteases Editors: A. B. Salleh et al., pp. 127-148

ISBN 1-60021-068-6 © 2006 Nova Science Publishers, Inc.

Chapter 8

MODIFIED LIPASES Bimo Ario Tejo, Kok Whye Cheong, Abu Bakar Salleh and Mahiran Basri ABSTRACT As demand grows, enzymes are being manipulated as an important catalyst to synthesize higher quality and specific products in both aqueous and non-aqueous media. This is due to their ability to exploit different substrates, high stability towards extreme conditions such as temperature, pH and tolerance to organic solvents. Other rationales include lipases procured chemo-, regio- and enantioslectivity, thus the products of lipase reactions have found wide applications in pharmaceuticals and fine-chemical industries. However, the natural properties of these enzymes created a stumbling block for their applications in harsh synthesis conditions. Modification needed to be carried out to modify the enzyme in order to have a better adaptability. Many techniques have been developed, e.g. modification using chemical modification, immobilization and protein engineering. Chemical modification has been the common approach as it offers troublefree method and gave exceptional properties to the modified lipases. Modification using aldehydes, polymers, anhydrides and imidoesters are aimed at creating a hydrophobic ‘blanket’ on the enzyme’s surface to promote solubility and higher activity in hydrophobic organic solvents. All the modified lipases have reported enhancement in activity, enantioselectivity, thermostability and stability towards organic solvents. Increasing degree of modification and hydrophobicity by using higher molecular weight modifiers, will generally increase the synthetic activity of the enzyme in organic solvents. Molecular dynamics studies have shown that by extensive alkylation, the open form of Candida rugosa lipase was stabilized, that may explain the increase of synthetic activity of alkylated lipase in organic solvent.

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INTRODUCTION Enzymes are natural catalysts made up of complex proteins, which have a great impact in our daily life. They act as catalysts in biochemical reactions and found numerous applications in food, feed, agriculture, paper, leather, and textiles industries, resulting in significant cost reductions (van Beilen and Li, 2002). As competitiveness among the industries grew stiffer, rapid technological developments are now stimulating the chemical, biotechnology, pharmaceuticals and fine-chemicals industries to use enzyme technology as the process are gaining new insights due to health, energy, raw materials and the environmental. Back in the 1980s, all the vast knowledge and studies of enzymology and its mechanism of reactions has been derived from studies of enzymes in aqueous solutions. The classic point of views pointed to the fact that enzymes work only in aqueous solutions but that is not the fact in today’s designation anymore. Enzymes have been employed for organic synthesis in organic solvents (Koskinen and Klibanov, 1996). As water is not the ideal reaction medium for most the organic processes, therefore, certain enzymatic conversions are better to be carried out in organic solvents for several reasons: 1. 2. 3. 4. 5. 6.

Organic substrates dissolve better in organic media. Alteration of substrate specificity. Enhanced enzyme stability. Enzymes are insoluble in organic media, thus, permits easy recovery and reuse. Product recovery from aqueous solutions is often difficult and expensive. Reduce undesirable side reactions, as water often takes part in reactions such as hydrolysis. 7. Thermodynamic equilibria of many processes are unfavourable in water. Example includes synthesis of esters from carboxylic acids and alcohols. Reactions in organic solvents would favour the formation of esters, where else, in water would favour the breaking down of esters into substrates. 8. Reduced microbial contamination. (Klibanov, 1989; Klibanov, 1988; Zaks and Klibanov, 1987; Klibanov, 1986)

However, the catalytic activity of enzymes in organic solvents is usually much lower than enzymes in an aqueous environment (Klibanov, 1997). Several reasons are attributed to masstransfer barriers for substrates (Klibanov, 2001), hydrophobic/hydrophilic balance on the protein surface (Ampon et al., 1993), greater tendency of stripping tightly bound water (which is essential for catalytic activity) from the enzyme molecules (Zaks and Klibanov, 1988), and reduced structural flexibility since organic solvents lack water’s ability to engage multiple hydrogen bonds (Affleck et al., 1992) and also lower dielectric constants which leads to stronger intra-protein electrostatic interactions (Klibanov, 2001). One of the most interesting and well-investigated class of enzymes in this particular field are lipases (triacylglycerol hydrolases, E.C. 3.1.1.3). Since lipases perform at the lipid-water interface, therefore, the presence of an organic phase is naturally essential to support the activation mechanism of these lipases. Lipases are among the popular ones and are frequently used because they catalyze various useful reactions, for instance hydrolysis, esterification, transesterrification and polyesterification reactions (Jääskeläinen et al., 1997), and act as

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chiral catalysts in the production of various fine-chemicals and intermediates (Berglund, 2001). The diverse functions and the enzyme specificity, both stereospecificity and regiospecificity make lipase one of the most important biocatalysts in biotechnological applications (Shaw, 2002).

AIMS AND ADVANTAGES OF MODIFICATION Why do we need to modify enzyme? Why chemical modification? Chemical modification is, above all, an important method to recognize essential side chains in active centre. In most cases, chemical modification of such side chains is followed by a decrease or an increase or changes in its catalytic activity. Chemical modifications hold some distinctive purposes (Pfleiderer, 1985): 1. To recognize essential side chains in the active centre. Functional amino-acid side chains with a high reactivity (so called ‘super-reactive groups’) are preferentially localized in the active centre of an enzyme. From the active centre, we understand those groups that are involved in the substrate binding and catalysis. 2. To distinguish between buried and exposed groups. To provide an idea about the topography of an enzyme/protein without the knowledge of the tertiary or quaternary structure. 3. For protein structural studies. Charged amino acids are located mostly on the surface of protein, and therefore could form interactions for stabilization of protein structure. Changes in the charge by modification will cause changes in protein conformations. 4. Chemical modification of essential groups can be used to introduce specific reagents as indicator groups, containing chromophoric, fluorescent or radical substituent. This would be helpful when physical measurements are carried out under conditions, which produce conformation changes, dissociation, et cetera. 5. Chemical modification of non-essential groups, mainly ε-amino a group of lysine plays an important role for biotechnology; the technical use of enzymes, especially for immobilization after covalent binding on insoluble matrices. The applicability of enzyme as industrial biocatalyst requires enzyme to withstand the ruthless environment. One of the main drawbacks for industrial utilization of biocatalysis is the inactivation of enzymes at elevated temperature and extreme pH. The efficient application of biocatalysts requires the availability of suitable enzymes with high activity and stability functioning under harsh conditions, desired substrate selectivity and high enantioselectivity. However, native enzymes which are without modification are limited and not specific to its application. These enzymes often need to be optimized to fulfil these requirements. Since the blooming of development in enzymology, the manipulation of the enzymes’ structures was seen as a new way to enhance its biochemical properties and stability, be it chemical modifications (Mine et al., 2001; Basri et al., 1998), protein engineering (Bornscheuer, 2002; Bornscheuer and Pohl, 2001; Harris and Craik, 1998) or by changing the reaction medium rather than the enzyme itself (Klibanov, 1989). Biochemical properties like activity, specificity and selectivity have been shown to be dependent on the choice of organic

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solvent and water activity (Berglund, 2001). Yet, on the other hand, modified enzymes with specific purpose are useful in certain fields. Therefore, many studies had been carried out in order to increase their activities, selectivity or solubility. Chemically modified enzymes was seen as a fruitful research in Basri and co-workers’ work in 1998 and Ampon et al. in 1991, as these lipases’ enantioselectivity and specificity were successfully enhanced. The applications of lipases or modified lipases in the industries (Sharma et al., 2001) as well as for synthetic purposes in organic media are now been widely reported (Tarahomjoo and Alemzadeh, 2003; Kiran and Divakar, 2001; Sereti et al., 1998; Basri et al., 1997); eg as biocatalysts for producing organic intermediates, specific chemicals and pharmaceuticals and as a biosensor.

TYPES OF MODIFICATION In the present day, enzymes and cells or organisms are widely been employed in biotechnology. Most mesophilic enzymes are marginally stabilized (Privalov, 1979), and after a long term of usage and storage, the activity of these commercial mesophilic enzymes would decrease and eventually lost their activity. Therefore, in today’s industrialized industry, a much more stable form of enzymes is much sought after. To obtain much more stable enzymes, many approaches have been made to stabilize commercial enzymes by various procedures using chemical and biological (genetic engineering) modifications (Mozhaev et al., 1988a). First and foremost, there are three different approaches for protein modifications, depending on the principles by which proteins are stabilized: 1. Amino acid residues of proteins are chemically modified, by specific chemicals such as polymers (de la Casa et al., 2001; Mine et al., 2001; Hernáiz et al., 1999; Zacchigna et al., 1998) aldehydes (Basri et al., 1998; Basri et al., 1997; Fujita et al., 1995; Ampon et al., 1991), imidoesters (Basri et al., 1992) and anhydrides (Song et al., 2005; Hosseinkhani et al., 2004; Bund and Singhal, 2002). 2. Proteins are bound or trapped physically onto supports or gels through chemical bonds. This immobilization technique has been applied to enzymes and cells or organisms, and has been widely employed in biotechnology (Mozhaev et al., 1988a; Klibanov, 1979). 3. A single or only a few amino acids in proteins are modified through substitution (protein engineering), especially through site-directed mutagenesis (Bornscheuer and Pohl, 2001), producing mutants of better physical and biochemical properties. In contrast to the physical and chemical practices, the candidate site for protein modification can be specifically modified at will.

CHEMICAL MODIFICATIONS OF LIPASE Different approaches have been applied to promote the reverse reaction of usual hydrolytic activity of lipases in microaqueous organic media. A strategy to improve the

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functionality of enzymes in organic solvents is by increasing the hydrophobicity of the enzyme’s surface, through chemical modification of the enzyme molecules. This can be done by attaching various hydrophobic compounds such as aldehydes or polymers which enables the enzyme to be soluble in a non-polar solution. Specifically, certain amino acid residues within the protein’s molecules, which play pivotal roles in protein conformation and catalysis, are derivatized with certain modifiers that may affect the enzymic properties. Originally one would expect that a distinct chemical reaction with a highly sensitive molecule like the enzyme is difficult to carry out, but under mild conditions (pH 5 – 9) and in aqueous solution most protein can react like a normal organic molecule characterised by a high molecular weight. This is attributed to the fact that the reactive groups of amino acid residues are limited. These residues act as nucleophiles and proton donors or acceptors, such as, carboxyl group (C-terminal, Asp and Glu), imidazolyl group (His), sulfhydryl group (Cys), amino group (N-terminal and Lys), hydroxyl group (Try, Thr and Ser), guanidinium group (Arg), indolyl group (Trp) and methylthio group (Met) (Nosoh and Sekiguchi, 1991). The first methods available to protein chemists to alter enzyme properties were in fact chemical modification. Previous to the discovery of site-directed mutagenesis, the chemical mutation of the serine protease subtilisin was accomplished by Polgar and Bender (Polgar and Bender, 1966) and Neet and Koshland (Neet and Koshland, 1966) who carried out the chemical conversion of the catalytic triad serine residue of subtilisin to a cysteine. If available, a reagent as specific as possible for a distinct functional group should be used. Absolutely specific are only a few reagents, such as imidoesters for amino groups, disulfides for thiols or tetranitromethane for tyrosines. Iodoacetic acid or iodoacetamide which was originally known as SH-blockers react forming the corresponding carboxymethylthioether of cysteine (Means and Feeney, 1971, 1995). The important aspect is to remove excessive reagents used. In many cases aliquots of the enzyme are denatured during the modification. Another important aspect that must be studied is the effect of modification on the properties of the modified enzyme. When the secondary, tertiary and quaternary structures are changed, then this can be the origin of activity changes. It is necessary to compare the native molecular weight, the circular dichroism spectra of the backbone or other characteristics of the native structure with those of the modified molecule (Pfleiderer, 1985). Chemical modification of proteins with polyethylene glycol (PEG) or PEGylation can be used to tailor molecular properties to particular applications. For biomedical applications, complexes of therapeutic proteins and PEG show reduced or eliminated immunoreactivity, prolonged clearance times and improved biostability. PEG-modified conjugates of the protein have been proven to reduce the immune reactions of human system towards therapeutic proteins of non-human origin. PEG-modified adenosine deaminase (PEG-ADA) is the first PEG-protein conjugate approved by the US Food and Drug Administration in 1991 (Levy et al., 1988). Though PEG-modified enzymes found applications in biomedical field, still, its major contributions would be nonaqueous enzymatic catalysis as nonaqueous media has numerous advantages compared to their use in conventional aqueous media. These enzymes found applications in surfactants, fine chemicals, textiles, pesticides and numerous synthesis industries (van Beilen and Zhi Li, 2002). Modification with PEG can also increase the solubility and activity of enzymes in organic solvents (Kodera et al., 1994), thus improving

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their potential for biotechnological applications. Some advantages of PEG-modified enzymes are (Inada et al., 1995): − − − −

increased catalysis of hydrophobic substrates; increased stereospecific synthesis in hydrophobic media; synthesis of compounds that are unstable in aqueous media; increased thermostability in both hydrophobic and aqueous media.

Another desired property of polyethylene glycol is its water-binding capacity that helps to keep a layer of water around the modified enzyme (Jene et al., 1997), which is known to be essential for catalytic activity in organic solvents (Zaks and Klibanov, 1988). To maintain an appreciable level of catalytic activity in nonaqueous media, certain number of bound water molecules is required to withhold its structure. Different enzymes showed vastly different need for the bound water in terms its amount (Zaks and Klibanov, 1988). Much work has been done so study the distinctiveness of PEG-modified lipase. In 1999, Hernáiz et al., conducted a systematic study of PEG-modified purified Candida rugosa lipase. The enzymatic heptyl oleate synthesis at 30ºC was found to increase due to the effect of modification; however the degree of modification did not change the synthetic activity. Nevertheless, at higher degree of modification, the derivative is more hydrophobic, therefore needs a smaller amount of water to be active. By using the PEG-lipases, high yields of esters were obtained in non-polar solvents (log P > 2). PEG-lipases showed very good solubility mainly in aromatic and chlorinated hydrocarbons. Gonzalez-Navarro and Braco (1998) attributed the enhanced activities to the modified lipases having attained the open activated conformation. The open structure of the lipases exposes the hydrophobic surfaces, which will interact with the hydrophobic substrates. Besides that, the active sites too, are uncovered and making them accessible to substrates. Else, Koops et al. (1999), treated lipases from Candida Antarctica with various hydrophobic groups, n-octanol, polyethylene glycol 2000 monomethyl ether (MPEG) and polyethylene glycol 400 mono-octyl ethyl (OPEG). The modified lipases were then tested for hydrolytic activity in water and transesterification activity in various organic solvents; oxylene, tert-butyl methyl ether, tert-butanol and 2-butanone. While the hydrolytic activity was slightly affected by modifications, the transesterification activities were strongly influenced. MPEG-modified lipases, being the least hydrophobic group among the three groups, showed 27-fold increase in transesterification activity. The more hydrophobic modifiers, OPEG and n-octanol showed very low activity and may lead to inactivation. The uniqueness of PEG-modified lipases does not stop here. Besides increasing in enzymatic activity in organic solvents, Zacchigna et al. in 1998 found that lipases from Candida rugosa modified using methoxy(polyethylene glycol)-p-nitrophenyl carbonate (NPC-mPEG) retained 98% of its hydrolytic activity of the non-modified enzyme. This mPEG-lipase is perfectly soluble in organic solvent and efficiently catalyze the esterification of lauric acid, with nearly 100 % of yield over six hours in toluene. In additional to the enhanced properties, mPEG-lipase displayed preference for the S-isomer over the R-isomer in the stereoselective resolution in racemic naproxen. In general, the solubility of the modified enzyme in organic solvent increases with increasing degree of modification. Inactivation of the enzyme however may occur during the

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133

derivatization, and this effect increases with the increasing degree of modification as reported by Ljunger et al. (1993) who studied the hydrolysis of benzoyltyrosine ethyl ester using polyethylene glycol modified α-chymotrypsin. They showed that while solubility of the modified enzyme in benzene increases with increasing degree of modification, catalytic activity decreases as the degree of modification is increased. A compromise with about half of the amino groups modified perhaps will provide useful applications of the modified enzymes. Derivatization of protein with PEG has several disadvantages. These include the toxicity of chemicals employed, the numerous steps necessary for the activation process, the low reactivity of intermediates, and the easy cleavage of the PEG-protein adduct (Ampon et al., 1991). Reductive alkylation of amino groups with aldehydes or ketones using cyanoborohydride as the reductant, is an alternative method which are simple and use less toxic reagents. Reductive alkylation is a convenient method to convert amino groups in proteins into their alkylamino derivatives. Many simple aldehydes and ketones react rapidly and reversibly with amino groups of protein to give a wide variety of alkyl substituents. The advantages of reductive alkylation include its technical simplicity, mild reaction conditions, high specificity for the α- and ε-amino groups, stability of the alkyl-lipase adducts and its comparative safety (Means and Feeney, 1995). Neither the initial adduct nor the Schiff base formed upon dehydration is very stable in dilute aqueous solution, but extensive modification of protein amino groups can be obtained by reduction of the Schiff base to a stable secondary amines. Step 1: protein-NH2 + RR’CO ' protein-NH-C(RR’)OH ' protein-N=C(RR’) Step 2: protein-N=C(RR’) NaCNBH protein-NH-HC(RR’) 3 As a rule, the reaction of proteins with aldehyde and the following reduction with NaCNBH3 is used. So far amino groups in proteins can be selectively and quantitatively converted into their alkyl derivatives at neutral pH under mild reaction conditions. Only the NH2 terminus and ε-amino groups are modified (Pfleiderer, 1985). In contrast to many other procedures for the modification of protein amino groups, reductive alkylation frequently has surprisingly little effect on the physico-chemical properties or biological activities of protein (Means and Feeney, 1971). At normal physiological pH, the typical alkylated ε-amino group will have essentially a unit positive charge, the same as the unmodified primary amino group. Reductive alkylation not only preserves the positive charges of protein amino groups, but, in contrast to other methods for their modification, does not change the approximate location of those charges. With small carbonyl compound like formaldehyde or acetaldehyde, reductive alkylation should have little or no effect on the distribution of charged groups and cause a minimal disturbance of existing electrostatic interactions (Pfleiderer, 1985). A wide variety of carbonyl compounds have been used for the reductive alkylation of proteins, be it as small as formaldehyde, to larger alkyl substituents, such as dodecylaldehyde. However, the mechanisms of reactions are usually similar to those with formaldehyde. By using formaldehyde as the carbonyl component, the reaction could proceeds rapidly giving ε-

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N,N-dimethyllysine as the main products. In the initial stage, monomethyllysine is formed, but its conversion to dimethyllysine is very fast as the monomethyllysine groups formed initially appear to be more reactive than unmodified amino groups and are converted to dimethyllysine groups under most conditions. With other aldehydes and ketones, reaction with second group of carbonyl molecule is greatly retarded such that predomantly monoalkyllysines are obtained (Means and Feeney, 1995; Means and Feeney, 1971). Introduction of large and bulky group, which are usually hydrophobic, via reductive alkylation prove to decrease the solubility of most proteins (Fretheim et al., 1979; Friedman et al., 1974). The modification of trypsin using dodecylaldehyde derives a completely insoluble derivative that is fully active after absorption to octyl-Sepharose (Wu and Means, 1981). Similar alteration using n-octanal has been used to stabilize trypsin’s interactions with an oil-water emulsion (Tsuji, 1990). Another technique to modify the amino group of amino acids is through amidination by imidoesters. This technique offer several advantages as the reaction conditions are relatively mild and the reaction is also specific for the ε-aminolysil and the α-group of the amino acids react in aqueous solution forming amidino groups in which the positive charge of the NH3+. The derivatised proteins retain its positive charge thus minimising the any changes in the ionic characteristics of the protein molecule (Makoff and Malcolm, 1981). The amidine linkages formed are stable over a wide range of experimental conditions (Hunter and Ludwig, 1962).

OUR WORK Chemical Modification with PEG PEGs have been attached to Candida rugosa lipase (CRL; E.C. 3.1.1.3) and PEGsmodified lipase had been used to synthesise some fatty esters (Basri et al., 1995). In this work, monomethoxypolyethylene glycols (PEG) of molecular masses 1900 and 5000 were activated using p-nitrophenyl chloroformate to form PEG-nitrophenyl carbonates, so called activated PEG (Scheme 1), with high yield (96-98%) and then was covalently attached to Candida rugosa lipase. These modified lipases exhibited specific ester synthesis activities on organic solvents compared with native lipase. The degree of activity enhancement depended on the size of activated PEG used and the degree of modification of the enzyme. Maximal activity was attained after exhaustive of modification (Table 1). The optimum esterification temperature (40°C) and preference of fatty acids as acyl donors of the modified lipase were very similar to those of the native enzyme. The modified lipase exhibited higher activity in non-polar solvents than in polar solvents (Figure 1), and showed higher temperature, solvent and storage stability then the native lipase (Figure 2-4).

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135

Table 1. Catalytic activities of PEG-lipases Activity (%) Syntheticb Hydrolytica

Modification PL1900 0 45 55 95 PL5000 48 58 63 95

100 3 2 1

100 300 415 912

5 5 4 3

130 189 399 643

a

Activity is expressed as % hydrolysis of olive oil emulsion of the native lipase. Activity is expressed as % of the native lipase activity. The ester synthesis is followed by the rate of disappearance of oleic acid from the reaction mixture containing propanol and oleic acid. (Source: Basri et al., 1995)

b

Specific activity (µmole min mg-1 protein)

2 1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0

He

e xan

gP (lo

.5) =3

nt rbo Ca

e

gP (lo ide r o chl tra

=3

)

B

gP (lo ne e z en

) =2 rm ofo lor h C

gP (lo Tri

) =2

ane eth o ro l h c

gP ( lo

/a) =n ine rid Py

gP (lo

m Di

.7) =0

i am rm lfo y eth

de

gP (lo

=-

Solvents

Figure 1: Catalytic activities of native-lipase and PEG-lipasesa in various organic solvents. Native lipase ( ) PL 1900 ( ) PL 5000 ( ) a The modification of enzyme preparations used were 45% and 48% for PL 1900 and PL 5000 respectively.

1)

O

O

O

OH

+

N+

O

O

Cl

O

N+

O

O-

methoxypolyethylene glycol

O-

O

O

p-nitrophenyl cloroformate

O

activated PEG

O N+

O

O-

O

+ H2N-Protein

pH 8.5, 28°C

O

O-

+

O Protein

O

O

O

activated PEG

Scheme 1: Modification of lipase with PEG, activated using p-nitrophenyl chloroformate.

N H

PEG-lipase

HO

N+ O

Modified Lipases

137

Residual activity (%)

120 100 80 60 40 20 0 20

30

40

50

60

70

80

Temperature (°C)

Figure 2:. Thermostability of PEG-lipases incubated for 1 h in benzene. NL (●), PL 1900 (▲), PL 5000 (■). (Source: Basri et al., 1995)

Residual activity (%)

120 100 80 60 40 20 0 0

5

10

15

20

25

Time (h)

Figure 3: Stability of PEG-lipases with respect to time at 40°C and 50°C. NL-40°C (●), NL-50°C (▲), PL-40°C (■), PL-50°C (♦). (Source: Basri et al., 1995)

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Residual activity (%)

120 100 80 60 40 20 0 0

5

10 Day

Figure 4: Stability of PEG-lipases incubated in benzene for 10 days at room temperature. NL (●), PL 1900 (▲), PL 5000 (■). (Source: Basri et al., 1995) R1 protein NH2 +

protein

C=O R2

protein

H

R1

N

C

H

R2

H

R1

N

C

O

OH

R2 -H2O

R1 protein

N

+H+ protein

C

-H+

H

R1

N

C R2

R2 Schiff base NaCNBH3

NaCNBH3

protein

H

R1

N

C R2

Scheme 2: Reductive Alkylation of Lipase.

H

Modified Lipases

139

Chemical Modification by Reductive Alkylation Salleh and coworkers (Salleh et al., 1990) have attached a variety of hydrophobic groups via reductive alkylation method with aldehydes of various chain length and steric hindrance to CRL (Scheme 2). The hydrolytic activity of modified enzyme was reduced which depended on the type of the aldehydes used for attachment. In the isooctane system, an increase in activity was observed with the short-chain aldehydes. For esterification activity, modified lipases showed higher activity than unmodified form with acetyl-lipase showed the highest activity among others (Figure 5).

Specific activity (µmole min mg-1 protein)

300

250

200

150

100

50

0 None

Acetaldehyde

Propionaldehyde

Benzaldehyde

Octaldehyde

Dodecyldehyde

PEG-lipase

Source of alkyl group Figure 5: Hydrolytic and synthetic activity of modified lipase. Aqueous system* (Hydrolysis) ( ) Isooctane** (Hydrolysis) ( ) Benzene*** (Synthesis) ( ) * Activity measured in an emulsified aqueous system containing equivolume olive oil and water, with polyvinylalcohol as an emulsifier. ** Activity measured in an assay system containing 50% vol olive oil, 30% vol water, and 20% vol isooctane. *** Activity of ester synthesis was based on the reaction of 1-propanol and oleic acid (6:1 mole ratio) in benzene.

The thermostability of native and alkylated lipase has been investigated at various temperatures. Enzymes derivatised with longer alkyl groups (octyl and dodecyl) were slightly more stable than those modified with shorter alkyl groups (Figure 6). Alkylated lipases were relatively more stable than native enzyme when they were incubated in benzene for various times (Figure 7) (Ampon et al., 1991).

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The effect of reductive alkylation of CRL on its selectivity has been examined (Basri et al., 1998). The enzyme could catalyze the enantioselective esterification of (R)-2-(4chlorophenoxy) propionic acid with n-tetradecanol. The optical purity, expressed as enantiomeric excess (ee) was higher for the alkylated lipase as compared to the native enzyme (Table 2). The percentage of conversions were increased with increasing degree of enzyme modification at longer reaction times than 24 hours (acetaldehyde) and 48 hours (octaldehyde) (Basri et al., 1998). Recently, experimental and theoretical approaches on alkylated lipase have been performed to study the structural properties of modified lipase in water and organic solvent. Far-ultraviolet circular dichroism (CD) of C. rugosa lipase in aqueous solvent showed that such chemical modifications at the enzyme surface caused a loss in secondary structure that is attributed to the enzyme unfolding which may be responsible for the decrease in activity of modified lipase in aqueous environment (Figure 8) (Rahman et al., 2004). An extensive molecular dynamics simulation has been carried out to study the effect of alkylation on the performance of C. rugosa lipase in carbon tetrachloride (CCl4). It has been shown that the open form of C. rugosa lipase is stabilized in the presence of hydrophobic solvent (Tejo et al., 2004). An extensive alkylation (95%) increased the stability of open form of C. rugosa lipase, thus may explain the increase of synthetic activity of alkylated lipase in organic solvent (unpublished result).

Residual esterolytic activity (%)

120

100

80

60

40

20

0 35

40

45

50

55

60

65

70

75

Temperature (ºC) Figure 6: Thermostability of native and alkylated lipase preparations. Samples of the enzyme preparation (3 mg ml-1) were incubated in 0.02 M phosphate buffer (pH 7) at various temperatures, and the residual esterolytic activity determined after 15 min. Native-lipase (♦), Ethyl-lipase (■), Propyllipase (▲), Benzyl-lipase (x), Octyl-lipase (+), Dodecyl-lipase (●). (Source: Ampon et al., 1991)

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141

120

Residual activity (%)

100

80

60

40

20

0 0

1

2

3

4

5

6

7

Day Figure 7: Stability of native, ethyl- and dodecyl-lipases in benzene incubated at room temperature for various time intervals. Native-lipase (♦), Ethyl-lipase (■), Dodecyl-lipase (▲). (Source: Ampon et al., 1991)

Table 2: Enantioselectivity of native and modified Candida rugosa lipase in esterification of 2-(4-chlorophenoxy) propionic acid with n-tetradecanol. Enzymes

Native lipase Acetyl-lipase

Octyl-lipase

Percent modification (%)

Enantiomeric excess

Conversion

E

0 38 58 70 43 63 95

0.02 0.03 0.03 0.09 0.03 0.66 0.76

0.04 0.1 0.12 0.21 0.25 0.28 0.51

3 1.64 1.62 2.16 1.24 7.95 13.62

Another enzyme that was modified by alkylation is trypsin (E.C. 3.4.4.4) (Ampon et al., 1993). The increment of surface hydrophobicity following the attachment of alkyl groups was correlated to the increment of its sugar esterification activity. These were dependent on the degree of modification and type of alkyl moieties bound to the enzyme molecules; however, excessive modification of the enzyme markedly reduced its esterification activity (Figure 9). Alkyl substitution of hydrogen will, however, necessarily increase both the bulk and hydrophobicity of the amino group and reduce its ability to form hydrogen bonds (Ampon et al., 1993).

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2000

molar residue ellipticity

1500 1000 500 0 200 -500

210

220

230

240

250

260

-1000 -1500 -2000 -2500

wavelength (nm)

Figure 8: Far-UV CD spectra of unmodified lipase, 49% lysines modified, 86% lysines modified and buffer (from bottom to uppermost, respectively). (Source: Rahman et al., 2004)

1800

Relative activity (%)

1350

900

450

0 0

20

40

60

80

100

Degree of modification (%) Figure 9: Relationship of the degree of trypsin modification using octaldehyde and acetaldehyde with their relative esterification activities determined using D-glucose and oleic acid as substrates in dimethylformamide. Ethyl-trypsin (■), Octyl-trypsin (▲).

Modified Lipases

143

Amidination of Lipase with Hydrophobic Imidoesters Another technique to introduce hydrophobic groups onto protein molecules is through amidination by imidoesters (Scheme 3). It had been shown that increasing the hyrophobicity of the imidoesters, increased the synthetic activity of the modified lipases while the hydrolytic activity was substantially decreased (Basri et al., 1992) (Figure 10). The hydrophobicity effect could also be enhanced by varying the degree of modification (Figure 11). As such, depending on our needs we can design the modification accordingly.

NH2 Protein - NH2

+

NH2

pH 8.5

Protein - NH - C - R

C-R

+

CH3OH

OCH3 Amidinated protein

Imidoester

Scheme 3: Modification of protein by amidination using imidoesters.

700 600

% activity

500 400 300 200 100 0 1

2

3

4

5

6

7

R group of imidoesters Figure 10: Effect of different hydrophobic group on the hydrolytica (●) and esterificationb (■) activity of modified lipase. The lipase was modified by different R groups of imidoesters as 1-unmodified, 2 – CH3 [Imidoester I], 3– (C6H5) [Imidoester II], 4– (C6H5)2 [Imidoester III], 5– CH3 (CH2)10 [Imidoester IV], 6 – (C6H5) (CH2)2 [Imidoester V], and 7– (C6H5)(CH2)3 [Imidoester VI]. a Activity was expressed as % of the hydrolysis of olive oil emulsion by the unmodified lipase. The rate of appearance of acid in the reaction mixture was followed by titration with 0.05 M NaOH. b The ester synthesis was followed by the rate of disappearance of oleic acid from the reaction mixture containing propanol and oleic acid as substrates in benzene. Titration of the remaining acid was with 0.05 M NaOH. Activity was expressed as % of the unmodified lipase activity.

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Modified enzymes have shown improved properties compared to its unmodified form, especially for reactions in hydrophobic media and at high temperature. Different modifiers and different degree of modification were shown to produce modified enzymes with different properties. Chemical modification thus, could be a method to change the enzyme properties tailored for different applications. However, besides being dependent on the degree of modification, the maximal activity of the modified enzyme was also shown to be dependent of the type of imidoesters used. To attain maximal activity, a lower degree of modification (63%) was needed with the more hydrophobic imidoester VI as compared to imidoester I (about 90%). As the degree of modification with imidoester VI was further increased beyond 63%, the modified enzyme showed a decrease in ester synthesis activity. This is parallel with the results obtained by Ljunger et al. (1993), whereas the activity of PEG-α-chymotrypsin dropped drastically with increasing degree of modification. This is due to the fact that imidoester is a hydrophobic compound and with the increasing degree of modification, the hydrophobicity of the enzyme’s surface would increase and prevent the accessibility of water molecule, which is known to be crucial for catalytic activity in organic solvents (Zaks and Klibanov, 1988). The native, catalytically active conformation of enzymes is maintained by various non-covalent interactions and water take parts in all of them (Arnold, 1988; Levinthal, 1986). Therefore, a small amount of water is vital in maintaining the protein conformation in nonaqueous systems. For this reason, the removal of water from within the structure would radically alter the protein conformation and destroy enzymatic activity. 14 12

Relative rate

10 8 6 4 2 0 0

48

63

75

91

100

% modification

Figure 11: Effect of increasing degree of modification on the esterfication activity. The imidoester used was with R – CH3. Relative rate is calculated by dividing the specific activity of the modified lipase at the indicated degree of modification with the specific activity of the unmodified lipase. (Source: Basri et al., 1992)

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145

REFERENCES Affleck, R., Haynes, C. A., and Clark, D. S., 1992. Solvent dielectric effects on protein dynamics. Proceedings of the National Academy of Science, USA, 89: 5167-5170. Ampon, K., Salleh, A. B., Salam, F., Wan Yunus, W. M. Z., Razak, C. N. A., and Basri, M., 1991. Reductive alkylation of lipase. Enzyme and Microbial Technology, 13: 597-601. Ampon, K., Salleh A. B., Basri, M., Yunus, W. M. Z., Razak, C. N. A., and Whitaker, J. R., 1993. Relationship of surface hydrophobicity to the sugar esterification activity of alkylated trypsin. Journal of Bioscience, 4 (2): 154-160. Arnold, F. H., 1988. Protein design for non-aqueous solvents. Protein Engineering, 2: 21-25. Basri, M., Ampon, K., Salleh, A. B., Yunus, W. M. Z., and Razak, C. N. A., 1992a. Catalytic activity of lipase modified with hydrophobic imidoester. Annals of New York Academy of Science, 672: 577-579. Basri, M., Ampon, K., Yunus, W. M. Z., Razak, C. N. A., and Salleh A. B., 1992. Amidination of lipase with hydrophobic imidoester. Journal of American Oil Chemists’ Society, 69 (6): 579-583. Basri, M., Ampon, K., Wan Yunus W. M. Z., Razak, C. N. A., and Salleh, A. B., 1995. Synthesis of fatty esters by polyethylene gycol-modified lipase. Journal of Chemical Technology and Biotechnology, 64: 10–16. Basri, M., Ampon, K., Wan Yunus, W. M. Z., Razak, C. N. A., and Salleh A. B., 1997. Enzymatic synthesis of fatty esters by alkylated lipase, Journal of Molecular Catalysis B: Enzymatic, 3: 171-176. Basri, M., Th’ng, B. L., Razak, C. N. A., and Salleh, A. B., 1998. Effect of reductive alkylation of Candida rugosa lipase on its enantioselective esterification reaction. Annals New York Academy of Science, 864: 192-197. Berglund, P., 2001. Controlling lipase enantioselectivity for organic synthesis. Biomolecular Engineering, 18: 13-22. Bornscheuer, U. T., and Pohl, M., 2001. Improved biocatalysts by directed evolution and rational protein design. Current Opinion in Chemical Biology, 5: 137-143. Bornscheuer, U. T., 2002. Methods to increase enantioselectivity of lipases and esterases. Current Opinion in Biotechnology, 13: 543-547. Bund, R. K., and Singhal, R. S., 2002. An alkali stable cellulase by chemical modification using maleic anhydride. Carbohydrate Polymers, 47: 137-141. de la Casa, R. M., Guisán, J. M., Sánchez-Montero, J. M., and Sinisterra, J. V., 2002. Modification of the activities of two different lipases from Candida rugosa with dextrans. Enzyme and Microbial Technology, 30: 30-40. Fretheim, K., Iwai, S., and Feeney, R. E., 1979. Extensive modification of protein amino groups by reductive addition of different sized substituents. International Journal of Peptide and Protein Research, 14 (5): 451-456. Friedman, M., Williams, L. D., and Masri, M. S., 1974. Reductive alkylation of proteins with aromatic aldehydes and sodium cyanoborohydride. International Journal of Peptide and Protein Research, 6 (3): 183-185. Fujita, Y., Hidaka, Y., and Noda, Y., 1995. Thermal stability of alkylated and hydroxyalkylated lysozymes. Thermochimica Acta, 253: 117-125.

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Gonzalez-Navarro, H., and Braco L., 1998. Lipase-enhanced activity in flavour ester reactions by trapping enzyme conformers in the presence of interfaces. Biotechnology and Bioengineering, 59: 122-127. Harris, J. L., and Craik, C. S., 1998. Engineering enzyme specificity. Current Opinion in Chemical Biology, 2: 127-132. Hernáiz, M. J., Sánchez-Montero, J. M., and Sinisterra J. V., 1999. Modification of purified lipases from Candida rugosa with polyethylene glycol: A systematic study. Enzyme and Microbial Technology, 24: 181-190. Hosseinkhani, S., Ranjbar, B., Naderi-Manesh, H., and Nemat-Gorgani, M., 2004. Chemical modification of glucose oxidase: Possible formation of molten globule-like intermediate structure. FEBS Letters, 561: 213-216. Hunter, M. J., and Ludwig, M. L., 1962. The reaction of imidoesters with proteins and related small molecules. Journal of American Chemical Society, 84 (18): 3491-3504. Inada, Y., Furukawa, M., Sasaki, H., Kodera, Y., Hiroto, M., Nishimura, H., and Matsushima, A., 1995. Biomedical and biotechnological applications of PEG- and PM-modified proteins. Trends in Biotechnology, 13: 86-91. Jääskeläinen, S., Linko, S., Raaska, T., Laaksonen, L., and Linko, Y., 1997. Molecular modelling of lipase-catalyzed polyester synthesis. Journal of Biotechnology, 52: 267-275. Jene, Q., Pearson, J. C., and Lowe, C. R., 1997. Surfactant modified enzyme: solubility and activity of surfactant-modified catalase in organic solvents. Enzyme and Microbial Technology, 20: 69-74. Kiran, K. R., and Divakar, S., 2001. Lipase catalyzed synthesis of organic acid esters of lactic acid in non-aqueous media. Journal of Biotechnology, 87: 109-121. Klibanov, A. M., 1979. Enzyme stabilization by immobilization. Analytical Biochemistry, 93: 1-25. Klibanov, A. M., 1986. Enzymes that work in organic solvents. Chemtech, 16: 354-359. Klibanov, A. M., 1988. Enzymatic catalysis in nonaqueous solvents. Journal of Biological Chemistry, 263: 3194-3201. Klibanov, A. M., 1988. The effect of water on enzyme action in organic media. Journal of Biological Chemistry, 263: 8017-8021. Klibanov, A. M., 1989. Enzymatic catalysis in anhydrous organic solvents. Trends in Biochemical Sciences, 14: 141-144. Klibanov, A. M., 1997. Why are enzymes less active in organic solvents than in water? Trends in Biotechnology, 15: 97-101. Klibanov, A. M., 2001. Improving enzymes by using them in organic solvents. Nature, 409: 241-246. Kodera, Y., Nishimura, H., Matsushima, A., Hiroto, M., and Inada, Y., 1994. Lipase made active in hydrophobic media by coupling with polyethylene glycol. Journal of American Oil Chemists’ Society, 71: 335-338. Koops, B. C., Verheji, H. M., Slotboom, A. J., and Egmond, M. R., 1999. Effect of chemical modification on the activity of lipases in organic solvents. Enzyme and Microbial Technology, 25: 622-631. Koskinen, A. M. P., and Klibanov A. M., 1996. Enzymatic Reactions in Organic Media. Blackie. London. pp: 20-50. Levinthal, C., 1986. Comment: “Dry” enzyme. Proteins: Structure, Function and Genetics, 1: 2-3.

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LIST OF CONTRIBUTORS Abu Bakar Salleh [email protected]

Azira Muhammad [email protected]

ABS is a Professor of Biochemistry, at the Department of Biochemistry, Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia. AM is a MS graduate and is now in US.

Bimo Ario Tejo [email protected]

BAT graduated with a PhD in 2004. He is now a postdoctoral fellow at Kansas State University.

Che Nyonya Abdul Razak

CNAR was an Associate Professor of Microbiology, at the Department of Biochemistry and Microbiology, Faculty of Science and Environmental Studies, Universiti Putra Malaysia. She is now happily retired and living a life of leisure. KWC is a graduate student pursuing his MS degree. FMS is a MS graduate and still looking for a job.

Kok Whye Cheong [email protected] Fairolniza Mohd Sharif [email protected] Thean Chor Leow [email protected] Mahiran Basri [email protected] Mohd Basyaruddin Abdul Rahman [email protected] Mohd Ropaning Sulong [email protected]

TCL is a PhD graduate. He is now a postdoctoral fellow in the Enzyme and Microbial Research group. MB is a Professor of Chemistry, at the Department of Chemistry, Faculty of Science, Universiti Putra Malaysia. MBAR is an Associate Professor of Chemistry, at the Department of Chemistry, Faculty of Science, Universiti Putra Malaysia. MRS was a MS graduate and is now working as a lecturer at the Universiti Industri Selangor.

150

A. B. Salleh, R. N. Z. R. A. Rahman and M. Basri

Noor Mona Md Yunus [email protected]

NMMY is a graduate student pursuing her MS degree.

Nor Azlina Ibrahim [email protected]

NAI is a PhD student in the Enzyme and Microbial Research group hoping to graduate this year.

Raja Noor Zaliha Raja Abd. Rahman [email protected] Sharul Nataqain Baharum [email protected]

RNZ is an Associate Professor of Microbiology, Department of Microbiology, Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia. SNB is a PhD graduate and is now working as lecturer at the Universiti Industri Selangor.

Siti Salhah Othman [email protected]

SSO is a PhD graduate. She is now a lecturer, at Kolej Universiti Islam Malaysia.

INDEX A access, 114, 117 accumulation, 65, 84 accuracy, 115 acetone, 47, 49, 65, 66, 67 acetonitrile, 70, 96, 97, 109 acid, 1, 2, 4, 5, 6, 7, 8, 9, 10, 11, 13, 14, 17, 21, 25, 26, 27, 29, 32, 35, 36, 37, 41, 46, 47, 51, 57, 64, 68, 74, 78, 84, 89, 91, 92, 100, 106, 107, 114, 118, 119, 120, 123, 129, 130, 131, 132, 135, 139, 140, 141, 142, 143, 146, 147 acrylic acid, 124 activated carbon, 120 activation, 9, 21, 23, 28, 63, 106, 116, 123, 128, 133 active site, 5, 8, 9, 16, 26, 27, 28, 29, 30, 38, 96, 112, 115, 116, 132, 147 acylation, 7, 27, 76, 96, 124, 147 adaptability, 127 additives, 2, 12, 14, 30, 52, 53 adenosine, 131, 147 adipose tissue, 3 adsorption, 111, 113, 114, 118, 119, 120, 121, 122, 124 affect, 63, 64, 69, 82, 89, 99, 112, 117, 119, 131 agar, 42, 49, 51, 55, 56, 69, 81, 98, 99 agent, 30, 31, 37, 73, 106, 115, 116, 121 aggregates, 2, 46, 48, 117 agriculture, 128 alanine, 4 alcohol(s), 2, 8, 10, 14, 65, 67, 123, 128 aldehydes, 127, 130, 131, 133, 134, 139, 145, 148 alkylation, 28, 127, 133, 134, 139, 140, 141, 145, 147, 148 alternative(s), 8, 14, 32, 33, 55, 58, 133 amines, 133 amino acids, 3, 5, 8, 23, 24, 25, 45, 51, 84, 101, 107, 112, 129, 130, 134

ammonia, 84 ammonium, 44, 45, 46, 47, 52, 66, 84, 100, 104 ammonium persulphate, 46 ammonium salts, 44 amylase, 31, 92, 116, 123 animals, 3, 23, 24, 26, 28, 33, 79 antibiotic, 80, 93 apoptosis, 36 aqueous solutions, 64, 128 arginine, 26 aromatic hydrocarbons, 99 arteries, 3 ASI, 16 Aspergillus terreus, 67, 69, 76 asymmetric synthesis, 21 ATP, 4 attachment, 113, 116, 139, 141 attacks, 10 attention, 13, 24, 41, 65, 97 autocatalysis, 25 autolysis, 23, 30 availability, 24, 129 awareness, 30

B Bacillus subtilis, 4, 5, 6, 15, 32, 39, 80 bacteria, 3, 6, 17, 20, 26, 29, 31, 33, 41, 42, 43, 45, 47, 49, 51, 58, 61, 63, 66, 69, 74, 81, 82, 83, 90, 91, 95, 98, 99 bacterial strains, 16, 59 bacterium, 18, 43, 51, 55, 60, 75, 76, 81, 99, 101, 104, 108, 109 barriers, 128 base catalysis, 29 benzene, 46, 64, 95, 98, 99, 102, 104, 133, 137, 138, 139, 141, 143 beverages, 13

152

Index

binding, 4, 9, 29, 34, 96, 108, 113, 115, 116, 118, 123, 129, 132 biocatalysts, vii, 1, 8, 12, 13, 14, 18, 20, 41, 70, 111, 112, 129, 130, 145 biodegradation, 14 biological activity, 112 biological processes, 23 biomass, 14, 36 biomedical applications, 131 bioremediation, 1, 96 biosynthesis, 2, 82 biotechnology, vii, 17, 19, 23, 24, 36, 58, 65, 69, 112, 128, 129, 130, 147 blocks, 9 blood, 12, 24, 31 bonds, 4, 23, 35, 98, 108, 114 brain, 3, 112, 123 branching, 3 breakdown, 3 breathing, 19 brevis, 79, 90 building blocks, 33

C calcium, 28, 30, 46, 48, 60, 80, 85, 89 candidates, 42, 68 carbohydrate, 24 carbon, 1, 11, 13, 41, 42, 43, 44, 49, 51, 52, 65, 68, 79, 83, 84, 85, 100, 101, 140 carbon atoms, 1 carbon tetrachloride, 140 carboxylic acids, 2, 128 carrier, 111, 113, 123 casein, 32, 51, 78, 80, 81, 84, 88, 97, 105, 106 catalyst, 7, 28, 35, 96, 111, 112, 114, 127, 147 catalytic activity, 15, 35, 46, 65, 68, 114, 119, 128, 129, 132, 133, 144 catalytic properties, 117 C-C, 109 cell, 13, 47, 57, 65, 71, 83, 86, 99, 101, 102, 103 cellulose, 117, 147 channels, 114, 115, 117 chemical bonds, 130 chemical interaction, 111 chemical properties, 9, 133 chicken, 79, 90 China, 45 chiral catalyst, 2, 129 chirality, 14 Chlamydia trachomatis, 5 chlorinated hydrocarbons, 132 chloroform, 73

cholesterol, 75, 124 chromatography, 11, 46, 47, 48, 49, 58, 66, 71, 72, 80, 86, 91, 97, 98, 104 chymotrypsin, 24, 26, 27, 28, 34, 35, 36, 89, 96, 98, 109, 124, 133, 144, 147 classes, 4, 7, 25, 26, 29, 80, 89 classification, 15, 26 cleaning, 1, 30 cleavage, 27, 29, 36, 133 clone, 32, 55, 56 cloning, 14, 15, 17, 18, 19, 20, 21, 55, 59, 60, 61, 74, 91 CO2, 124 coagulating enzyme, 32 cobalt, 29 cocoa butter, 1, 2, 13, 18 collagen, 33 commitment, vii commodity, vii compatibility, 37, 117 competitiveness, 128 components, 8, 13, 30, 31 composites, 113 composition, 8, 43, 91, 116, 118, 119 compounds, vii, 2, 8, 14, 28, 29, 31, 33, 44, 45, 112, 115, 131, 132, 133 concentration, 43, 46, 55, 57, 63, 64, 65, 67, 71, 84, 85, 87, 89, 104, 112, 116 condensation, 35 consciousness, vii consensus, 6, 106 conservation, 30 consumers, 32 contamination, 42, 112, 128 control, 13, 15, 34, 48, 51, 52, 55, 71, 99, 102, 106, 107 conversion, 36, 131, 134, 147 copper, 29 corn, 41, 44, 51, 52, 66, 84, 100 correlation, 83 costs, 8 cotton, 14 coupling, 112, 114, 115, 117, 146 covalent bond, 29 crude oil, 14 crystallization, 92 cultivation, 15, 43, 44, 55, 59, 66, 86, 99 culture, 17, 44, 46, 47, 52, 55, 65, 70, 71, 77, 83, 86, 99, 101, 102 culture media, 99 cutin, 2 cycles, 78, 115

Index

D database, 26 decomposition, 12 deficiency, 147 degradation, 12, 21, 23, 33, 34, 38, 96 dehydration, 133 demand, 3, 32, 127 denaturation, 3, 14, 42, 51, 77, 86, 112, 116 Denmark, 24 density, 36, 114, 117 dentures, 30 deposition, 3, 124 depression, 82 derivatives, 1, 3, 10, 122, 133 detection, 69 detergents, 1, 12, 14, 30, 31, 32, 37, 47, 78, 98, 147 dielectric constant, 128 diesel engines, 2 diffusion, 42, 113, 117, 119 diffusion rates, 42 digestion, 3, 12 dimerization, 29 dimethylformamide, 65, 142 dissociation, 129 distribution, 117, 133 diversity, 24, 79 division, 3 DMF, 65, 97 donors, 35, 131, 134 drugs, 1 drying, 13, 87 duration, 44

E earth, 68 effluent, 24, 33, 42, 49, 50 elastin, 33 elastomers, 124 electrophoresis, 47 employment, 66 enantiomers, 17 encoding, 5, 45, 55, 86, 92 endonuclease, 57 England, 18, 147 entrepreneurs, 33 environment, 8, 12, 14, 31, 41, 95, 112, 128, 129, 140 environmental awareness, 31 enzymatic activity, 96, 113, 115, 132, 144, 148 enzyme immobilization, 112, 113, 114, 117, 119

153

enzymes, vii, 1, 2, 3, 4, 5, 6, 7, 12, 14, 15, 16, 18, 23, 26, 27, 28, 29, 30, 31, 34, 35, 36, 41, 42, 47, 59, 63, 64, 65, 67, 68, 70, 75, 77, 78, 79, 80, 86, 87, 95, 96, 97, 104, 108, 109, 111, 112, 115, 116, 117, 118, 123, 127, 128, 129, 130, 131, 132, 133, 144, 146 epoxy groups, 116 equilibrium, 8, 34, 95, 96, 117, 119 ester, 1, 2, 7, 8, 9, 10, 11, 17, 18, 19, 29, 35, 49, 67, 74, 89, 122, 123, 133, 134, 135, 139, 143, 144, 146 ester bonds, 7, 8, 9, 11 ethanol, 35, 65, 67 ethyl acetate, 104 ethylene, 65, 124, 147 ethylene glycol, 65, 147 ethylene oxide, 124 eukaryotic cell, 29 evidence, 96 evolution, 80, 145 excretion, 86 experimental condition, 116, 134 exposure, 11, 41, 51, 105, 112 expression, 15, 18, 19, 21, 41, 45, 48, 55, 57, 58, 59, 60, 90, 91, 92, 107, 108 extraction, 46, 66

F family, 4, 5, 6, 28, 89 fat, 1, 3, 8, 13, 14, 18, 19, 34, 41 fatty acids, 1, 7, 8, 9, 11, 12, 13, 14, 15, 46, 75, 124, 134 fermentation, 16, 34, 36, 37, 44, 82, 101, 125 fibers, 115 filtration, 46, 47, 48, 66, 80, 86 fish, 24, 46, 75, 124 fish oil, 46, 75, 124 fishing, 97 flavor, 1, 2, 12, 13 flexibility, 128 fluorescence, 43, 55 food, 1, 2, 3, 12, 13, 23, 24, 67, 128 food additives, 12 food industry, 2, 13, 23, 24 food production, 67 formaldehyde, 133 fructose, 51, 100, 101 fuel, 2, 13, 16, 124 fungus, 13

154

Index

G gastric mucosa, 25 gel, 11, 46, 47, 48, 66, 80, 86, 114, 115, 120, 124, 125 gene, 15, 16, 18, 19, 21, 23, 32, 45, 48, 55, 57, 74, 77, 86, 91, 92, 106, 108 gene expression, 23 genes, 41, 45, 106 genetics, 21 Germany, 19, 38 germination, 23 glucose, 43, 44, 51, 66, 79, 101, 142, 146 glucose oxidase, 146 glucoside, 14 glutamate, 8 glutamic acid, 29 glutathione, 45 glycerol, 1, 7, 9, 11, 13, 14 glycine, 4, 32 glycol, 65, 117, 118, 131, 132, 133, 146, 147, 148 grass, 14, 31 grouping, 4 groups, 3, 9, 10, 11, 12, 24, 25, 26, 31, 32, 35, 116, 117, 129, 131, 132, 133, 134, 139, 141, 143, 145 growth, 16, 41, 42, 43, 44, 52, 59, 66, 69, 70, 79, 80, 82, 83, 84, 85, 88, 99, 101, 102, 116 growth rate, 44, 84 growth temperature, 42, 80, 82, 88 Guinea, 18, 60

H hair follicle, 33 half-life, 46, 47, 58, 72, 77, 78, 80, 105 health, 33, 128 heat, 6, 32, 42, 43, 44, 49, 78, 79, 86, 88, 90, 92, 98, 116, 119, 121 heating, 11, 121 heptane, 67 herbicide, 14, 21 hexane, 11, 46, 64, 65, 67, 70, 71, 96, 102, 104, 109, 119 higher quality, 127 histidine, 8, 28, 29, 101 HIV, 29 homogeneity, 3, 45, 46, 47, 48, 66, 71, 86, 104 host, 41, 45, 55, 107 human immunodeficiency virus, 36 hydrogels, 111, 118 hydrogen, 10, 33, 64, 128, 141 hydrogen bonds, 64, 128, 141

hydrogen peroxide, 10 hydrolysis, 1, 2, 7, 8, 9, 10, 11, 12, 18, 19, 20, 21, 23, 25, 33, 34, 42, 47, 49, 68, 74, 87, 97, 105, 123, 128, 133, 135, 143 hydrophilicity, 111, 119 hydrophobicity, 64, 117, 118, 127, 131, 141, 143, 144, 145 hydroxide, 119 hydroxyl, 27, 28, 115, 123, 131 hydroxyl groups, 115, 123

I ibuprofen, 16 ideas, 36 identification, 31, 75, 81 immobilization, 39, 111, 112, 113, 114, 115, 116, 117, 118, 119, 120, 121, 122, 123, 124, 125, 127, 129, 130, 146 immobilized enzymes, 111, 112, 114, 115, 116, 117, 121, 123 immune reaction, 131 immunoreactivity, 131 impurities, 14, 66 in vitro, 92 inclusion, 114, 115 incubation time, 101 India, 13, 32 Indonesia, 44 inducer, 55, 57, 85 induction, 55, 57, 58, 82 induction time, 55, 57 industrial processing, 41 industry, vii, 1, 3, 7, 14, 20, 23, 24, 31, 32, 33, 34, 41, 67, 68, 77, 96, 97, 130 influence, 116 infrared spectroscopy, 15 inhibition, 14, 43, 46, 51, 69, 101, 106, 112, 116 inhibitor, 28, 29, 30, 35, 37, 38, 89, 96, 106 injury, 36 inoculum, 49, 70, 99, 100 insertion, 4 insight, 43, 96 instability, 95 intensity, 55, 86 interaction, 48, 66, 70, 71, 95, 97, 98, 104, 113, 114, 117, 118 interactions, 118, 128, 129, 133, 134, 144 interest, 2, 31, 77, 96, 111 interface, 1, 9, 12, 36, 128 iodine, 11 ions, 29, 46, 47, 48, 49, 51, 52, 54, 58, 68, 69, 70, 73, 80, 89, 101, 121

Index Islam, 150 isolation, vii, 41, 42, 64, 69, 78, 97 isomers, 103

J Japan, 13, 31

K keratin, 33 ketones, 133 kinetic parameters, 15, 59 kinetics, 20, 123 knowledge, 3, 35, 77, 114, 128, 129

L lactate dehydrogenase, 57 lactic acid, 146 lactose, 51, 79, 100, 101 land, 24 lead, 34, 41, 105, 112, 132 leisure, 149 leucine, 29 ligands, 29 limitation, 44 linkage, 116 lipases, vii, 1, 2, 3, 4, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 41, 42, 43, 44, 47, 48, 49, 51, 59, 60, 63, 65, 67, 68, 69, 71, 72, 73, 75, 111, 112, 118, 120, 121, 122, 123, 124, 125, 127, 128, 130, 132, 134, 135, 137, 138, 139, 141, 143, 145, 146, 147 lipid metabolism, 3 lipids, 2, 3, 12, 14, 19, 42 lipolysis, 9 liver, 36 location, 133 low temperatures, 31 lubricants, 2, 14 lysine, 26, 27, 28, 129 lysis, 55, 98

M Malaysia, vii, 49, 91, 95, 98, 108, 124, 149, 150 maltose, 51, 101 manganese, 89 manipulation, 45, 79, 129 mannitol, 51

155

manufacturing, 13 mapping, 108 marine environment, 38 market, 24, 31, 32, 77 market share, 77 mass, 4, 12, 49, 107, 128 mass spectrometry, 49 matrix, 38, 98, 113, 114, 115, 117, 122, 125 matrix metalloproteinase, 38 measurement, 21, 116 measures, 67 meat, 24 media, 19, 35, 55, 63, 64, 66, 70, 74, 75, 76, 81, 82, 83, 95, 96, 99, 100, 101, 102, 127, 128, 130, 131, 132, 144, 146, 147 melting, 42 membranes, 3, 16, 122, 123 memory, 96 metabolism, 3 metals, 14 methanol, 13, 46, 49, 64, 65, 67, 73, 97 methodology, 16, 44 mice, 36 microorganism, 1, 3, 44, 49, 81, 82, 97, 98, 99 microspheres, 122 migration, 10 milk, 3, 8, 13, 15, 19, 25, 32, 65, 81, 98, 99 mixing, 114 mobility, 34 mode, 29 modeling, 124 moisture, 7, 34 mole, 57, 139 molecular dynamics, 140 molecular mass, 4, 6, 46, 49, 55, 66, 71, 88, 97, 98, 107, 134 molecular weight, 21, 28, 32, 46, 47, 105, 118, 127, 131 molecules, 29, 46, 64, 96, 97, 106, 115, 118, 119, 121, 128, 131, 132, 141, 143, 146 monolayer, 117 monomers, 118 morphology, 117, 124 movement, 115 MRS, 149 mutagenesis, 14, 130, 131 mutant, 35 mutation, 131

N NaCl, 43 nanocomposites, 119

156

Index

nanoparticles, 116, 123 natural polymers, 117 needs, 132, 143 Netherlands, 24 network, 34, 38, 115, 117 nickel, 29 nitrogen, 41, 44, 51, 52, 53, 65, 79, 83, 84, 100, 101 nucleophiles, 131 nucleotides, 95

O octane, 67 oil, vii, 2, 7, 8, 9, 11, 12, 14, 15, 17, 18, 20, 41, 42, 43, 44, 46, 47, 48, 49, 50, 53, 54, 55, 57, 58, 59, 65, 66, 68, 70, 73, 81, 124, 134, 135, 139, 143 oils, vii, 11, 16, 18, 47, 51, 53, 65, 68 operator, 9, 86 optical density, 99 optimization, 16, 50, 55, 65, 75, 117 organic matter, 42 organic solvents, 1, 7, 10, 12, 18, 22, 34, 35, 42, 63, 64, 65, 67, 70, 71, 73, 75, 76, 80, 95, 96, 97, 102, 103, 104, 106, 107, 108, 109, 121, 127, 128, 131, 132, 134, 135, 144, 146, 147, 148 organism, 3, 29, 65, 82, 99 orientation, 117 oxygen, 14, 29, 83, 99

P palm oil, vii, 3, 8, 13, 18, 19, 21, 42, 49, 50, 53, 68, 69, 73, 74, 82 pancreas, 24 parameter, 30 particles, 2, 14, 18 partition, 64 pasteurization, 32 PCR, 55 pepsin, 24, 28, 32 peptidase, 25, 35, 37 peptides, 7, 23, 34, 35, 36, 37, 97 permit, 14 pH, 1, 8, 12, 14, 27, 28, 29, 30, 33, 41, 42, 43, 44, 46, 47, 48, 49, 58, 59, 64, 66, 67, 68, 70, 72, 73, 77, 78, 80, 81, 82, 83, 87, 97, 98, 99, 100, 101, 105, 106, 112, 113, 114, 115, 116, 127, 129, 131, 133, 140 pharmaceuticals, 12, 14, 127, 128, 130 phenylalanine, 29, 89, 114, 122 phospholipids, 2, 19 phylogenetic tree, 3

physical interaction, 112 physical properties, 13, 112 physicochemical properties, 91 pigs, 24, 32 pitch, 13 plants, 3, 23, 24, 26, 28, 79 plasmid, 48, 49, 55, 58 PM, 146 polarity, 64, 70, 96, 118 pollutants, 96 pollution, 31, 33 poly(2-hydroxyethyl methacrylate), 122 polyacrylamide, 47, 117 polymerase, 106 polymerase chain reaction, 106 polymers, 23, 111, 113, 117, 118, 127, 130, 131 polypeptide, 5, 106, 107 polypropylene, 124 polystyrene, 117 polyurethane, 12, 21 polyvinylalcohol, 139 poor, 32, 63 porosity, 115, 117 potassium, 46 precipitation, 46, 47, 66, 104 preference, 2, 11, 47, 96, 132, 134 preparation, 17, 34, 66, 116, 124, 125, 140 pressure, 8, 33, 68, 117 principle, 118 producers, 4, 24, 25, 42, 44, 49, 50, 63, 69, 70, 71, 78, 79, 95, 97 production, 2, 12, 13, 14, 15, 16, 17, 18, 24, 27, 31, 32, 33, 34, 35, 37, 41, 43, 44, 45, 49, 51, 52, 53, 54, 55, 58, 60, 63, 65, 66, 69, 70, 74, 75, 77, 78, 79, 82, 83, 84, 85, 92, 95, 98, 99, 100, 101, 102, 108, 109, 116, 121, 124, 129, 148 productivity, 13, 43 promoter, 45, 48, 86 propane, 29 propylene, 65 protease inhibitors, 30 protective coating, 2 protein crystallization, 51 protein design, 145 proteinase, 25, 26, 28, 37, 39, 90, 91, 92 proteins, 23, 33, 34, 42, 45, 64, 66, 70, 86, 88, 130, 131, 133, 134, 145, 146, 148 proteolysis, 32, 34, 106 proteolytic enzyme, 90, 97, 98, 102, 104, 108 protocol, 48 Pseudomonas aeruginosa, 4, 5, 6, 19, 21, 44, 64, 65, 66, 67, 73, 75, 76, 95, 97, 98, 99, 102, 104, 106, 108, 109

Index purification, vii, 13, 15, 17, 20, 41, 46, 48, 51, 58, 59, 60, 61, 66, 69, 70, 71, 74, 75, 76, 80, 86, 90, 91, 92, 95, 104, 107, 108, 109, 147

R race, 132 range, vii, 1, 2, 4, 27, 29, 30, 31, 42, 53, 58, 67, 70, 72, 73, 77, 80, 82, 84, 87, 98, 99, 105, 134 rape, 3 raw materials, 128 reaction mechanism, 26, 36 reaction medium, 128, 129 reaction rate, 117 reaction time, 140 reactive groups, 129, 131 reading, 6, 45, 55, 106, 107 reagents, 30, 115, 129, 131, 133 recombinant DNA, 32, 45 recovery, 46, 47, 48, 49, 66, 86, 128 recycling, 112 reduction, 18, 42, 51, 84, 99, 133 refining, 14, 18 regioselectivity, 64 regulation, 33 relationship, 4, 147 relationships, 38 relevance, 3 renin, 24, 28, 32 repression, 43 residues, 4, 5, 6, 9, 16, 26, 27, 29, 33, 36, 89, 92, 106, 107, 130, 131 resistance, 64, 116 resolution, 1, 20, 21, 35, 36, 37, 39, 96, 120, 132 resources, vii retention, 34, 93 rheology, 34 rice, 43, 44 risk, 42 room temperature, 8, 138, 141 rubber, 115

S safety, 70, 133 sales, 12, 23, 24, 30, 79 salts, 80 sample, 39, 69, 93 sampling, 81 saturated fat, 46 saturated fatty acids, 46 saturation, 118

157

search, vii, 32, 69, 96, 116, 119 secrete, 82, 99 secretion, 4, 23, 55, 91 sediment, 39 seed, 18 selectivity, 17, 20, 37, 42, 112, 117, 123, 129, 140 sensitivity, 42, 98 separation, 8, 46, 111 sequencing, 15, 21, 59, 91, 108 series, 16 serum, 3, 57, 88, 104 serum albumin, 57, 104 shortage, 32 signal peptide, 45, 55, 57 silica, 11, 113, 115, 116, 117, 120, 123, 124, 125 similarity, 4, 6 simulation, 124, 140 sites, 26, 49, 50, 81, 96 skin, 18, 33, 37 sludge, 14 sodium, 31, 33, 47, 52, 84, 85, 100, 101, 119, 145 soils, 98 sol-gel, 115 solid phase, 112, 148 solubility, 34, 41, 42, 58, 63, 96, 119, 127, 130, 131, 132, 134, 146 solvent molecules, 96 solvents, 58, 63, 64, 65, 67, 70, 95, 96, 97, 98, 102, 104, 106, 127, 128, 132, 134, 145, 146, 148 species, 2, 65, 67, 79, 91, 97, 106 specific surface, 115, 116, 119 specificity, 1, 2, 7, 9, 11, 12, 13, 14, 19, 24, 26, 27, 32, 35, 46, 58, 63, 64, 73, 80, 89, 95, 96, 117, 120, 121, 128, 129, 133, 146 spectrum, 1 speed, 33 spleen, 3 spore, 23, 99 stability, 1, 8, 28, 30, 31, 41, 42, 48, 58, 59, 63, 64, 65, 66, 67, 68, 70, 72, 73, 80, 86, 87, 97, 98, 102, 103, 104, 105, 106, 107, 108, 112, 115, 116, 117, 119, 120, 121, 127, 128, 129, 133, 134, 140, 145, 147, 148 stabilization, 67, 90, 112, 124, 129, 146 stages, 3 starch, 51, 66, 79, 84, 99, 100, 101, 123 stereospecificity, 13, 129 steroids, 2 stock, 54 stomach, 25, 32 storage, 32, 115, 121, 130, 134 strain, 3, 11, 15, 21, 24, 38, 39, 41, 42, 43, 45, 46, 48, 49, 51, 52, 53, 54, 55, 59, 60, 61, 63, 64, 66,

158

Index

69, 70, 71, 72, 73, 74, 77, 78, 80, 81, 82, 90, 91, 92, 95, 97, 98, 99, 100, 101, 102, 103, 104, 105, 106, 107, 108, 109 strength, 64 strong interaction, 45 structural gene, 45 students, vii substitutes, 13 substitution, 130, 141 substrates, 1, 2, 8, 11, 12, 19, 21, 27, 34, 35, 41, 42, 44, 46, 47, 53, 54, 63, 64, 67, 68, 70, 88, 96, 106, 117, 120, 127, 128, 132, 142, 143 sucrose, 51, 79, 84, 101 sugar, 141, 145 suppliers, 31 surface area, 9, 112, 117, 118, 119 surface properties, 115 surfactant, 14, 31, 119, 146 surprise, 64 susceptibility, 79, 106 Sweden, 71 swelling, 33 synergistic effect, 100 synthesis, 1, 2, 7, 8, 12, 14, 15, 16, 20, 22, 34, 35, 36, 37, 39, 52, 67, 95, 97, 115, 122, 127, 128, 131, 132, 134, 135, 139, 143, 144, 145, 146, 147 systems, 13, 24, 65, 113, 144

T technological developments, 128 technology, 15, 32, 45, 128, 148 temperature, 1, 24, 30, 31, 33, 41, 42, 45, 46, 47, 48, 51, 55, 58, 59, 68, 70, 72, 73, 78, 80, 82, 88, 92, 97, 99, 101, 102, 105, 113, 114, 115, 127, 129, 134, 144 textiles, 128, 131 thermal activation, 60 thermal stability, 32, 48, 64, 74, 77, 96 thermodynamic equilibrium, 8 thermostability, 42, 60, 64, 72, 77, 78, 86, 89, 105, 111, 116, 127, 132, 139 thrombin, 26 time, vii, 11, 24, 31, 34, 41, 44, 48, 49, 55, 64, 79, 116, 137, 141 tissue, 3 toluene, 64, 65, 73, 74, 75, 95, 98, 99, 102, 104, 132 toxicity, 14, 133 trace elements, 55 trade, 30 transcription, 55 transesterification, 1, 7, 13, 18, 19, 21, 74, 132 transformation, 9, 16

transformations, 10 transition, 27, 68 transition metal, 68 translation, 23, 55 translocation, 45 transport, 14 trend, 114 triacylglycerides, 1 triglycerides, 7, 9, 11, 12, 13, 20, 42, 46, 47, 49, 73 trypsin, 24, 26, 27, 28, 33, 89, 125, 134, 141, 142, 145, 148 tryptophan, 25, 30 tumor, 36 tumor necrosis factor, 36 tyrosine, 25, 104

U UN, 107 United Kingdom, 21 United States, 24 urea, 106 UV, 43, 142

V values, vii, 13, 70, 95, 97, 102, 105, 106 variable(s), 19, 68 variation, 83, 116 vector, 45, 55, 58, 107 velocity, 9 versatility, 12 vessels, 8 viscosity, 70

W waste water, 12, 14, 18, 97 water, 1, 7, 8, 9, 10, 12, 13, 14, 16, 18, 20, 22, 27, 28, 29, 34, 42, 46, 63, 64, 65, 67, 70, 74, 75, 76, 96, 97, 109, 116, 119, 121, 124, 128, 130, 132, 134, 139, 140, 144, 146, 147 wheat, 35 wood, 13, 34, 98 work, vii, 34, 75, 111, 115, 116, 117, 119, 120, 128, 130, 132, 134, 146 workers, 33, 118

X XRD, 119

Index

Y yeast, 15, 42, 44, 51, 52, 53, 55, 66, 70, 100 yield, 7, 32, 46, 48, 51, 52, 65, 66, 67, 75, 78, 82, 83, 84, 85, 86, 99, 100, 104, 123, 132, 134

159

Z zeolites, 39, 120 zinc, 29