ALKALITHERMOPHILIC LIPASES FROM THERMOSYNTROPHA LIPOLYTICA

by

MOH’D A. SALAMEH

(Under the Direction of JUERGEN K. W. WIEGEL)

ABSTRACT

Thermosyntropha lipolytica DSM 11003 is an anaerobic thermophilic alkalitolerant

bacterium which grows syntrophically with a methanogen on lipids utilizing only the fatty

acid moieties, but not the glycerol. Since no lipases from thermophilic anaerobes have

been analyzed, there is the possibility to find enzymes with different stereo and

substrate specificity not observed among the Gram negative aerobic ones. The

objective of this study was to characterize the lipases that are produced by this

bacterium with the intent of producing commercially viable enzymes for use in high

temperature, high pH-value applications, and various organic synthesis reactions such

as structured lipids, remodeling acyl-alcohols and the resolution of racemates.

Two enzymes, termed LipA and LipB, were purified from the culture supernatant

to gel electrophoretic purity by ammonium sulfate precipitation and column

chromatography using Octyl Sepharose fast flow. The apparent molecular weight of

LipA and LipB determined by SDS-PAGE were 50 and 57 kDa, respectively. The

temperature optima of purified LipA and LipB was 96 °C, this is the highest among all

known lipases so far. The pH25°C optima of LipA and LipB were 9.4 and 9.6, respectively.

They are among the most thermostable lipases with a half life of 24 h at 75 °C. Both

enzymes are true lipases that are interfacially activated by the presence of insoluble

substrates. They prefer glycerides with long chain fatty acids (C12 to C18), and prefer to

hydrolyse ester bonds at the primary position.

In addition, the effect of different detergents on enzymes activity and stability was

investigated. SDS was found to have the highest impact on both lipases by enhancing

the activity of both LipA and LipB by approximately 9 folds. Assay analyses using the

serine inhibitor E600 with increasing concentration of SDS and Tween 20 strongly

suggest that SDS and Tween 20 promote conformational changes by binding to the lid

domain and/or active site pocket so that the site becomes accessible to the substrate.

The purified native lipases showed a strong tendency to form catalytically active

oligomers as observed by gel filtration chromatography. This property might be a major

contributor to their high thermostability.

Finally, the ability of both lipases to conduct “reverse” synthesis reactions was

investigated. The maximum catalytic activity was measured at 85°C in isooctane. Octyl

oleate and lauryl oleate were the highest conversion products of the esterefication

reactions. In addition, LipA and LipB effectively catalyzed the synthesis of

diacylglycerols, in particular 1,3-dioleoyl glycerol.

INDEX WORDS: Thermosyntropha lipolytica, Alkalithermophilic, Lipases, gel filtration,

protein oligomerization, lipase purification, thermostability, thermal activity, p-nitrophenyl laurate, SDS, critical micelle concentration, E600, Tween 20, detergents, regiospecificity, stereospecifity, fatty acid esters, fatty acid alcohols, organic synthesis, esterification, triacyl glycerols, diacyl glycerols, oleic acid.

ALKALITHERMOPHILIC LIPASES FROM THERMOSYNTROPHA LIPOLYTICA

by

MOH’D A. SALAMEH

B.S. Yarmouk University, Jordan, 1995

M.S. Middle East Technical University, Turkey, 1999

A Dissertation Submitted to the Graduate Faculty of The University of Georgia in Partial

Fulfillment of the Requirements for the Degree

DOCTOR OF PHILOSOPHY

ATHENS, GEORGIA

2006

© 2006

Moh’d A. Salameh

All Rights Reserved

ALKALITHERMOPHILIC LIPASES FROM THERMOSYNTROPHA LIPOLYTICA

by

Moh’d A. Salameh

Major Professor: Juergen Wiegel

Committee: Michael W. Adams Anna Karls Robert Maier William B. Whitman

Electronic Version Approved: Maureen Grasso Dean of the Graduate School The University of Georgia August 2006

ACKNOWLEDGEMENTS

I would like to express my greatest appreciation to my supervisor Prof. Juergen

Wiegel. I would like to thank him for his enormous support and help, and for always

being there for me over the past six years of my graduate studies at The University of

Georgia. I am forever grateful for his willingness to share his extraordinary knowledge

with me, and his encouragement in my steps to become an independent researcher.

Many thanks and appreciation to my great committee members, Dr. Michael Adams, Dr.

Robert Maier, Dr. Anna Karls and Dr. Barny Whitman for their enthusiasm and

numerous helpful discussions, comments and suggestions which were extremely helpful.

I also want to thank the Microbiology Department for providing a teaching assistantship

for the past 6 years of my graduate school. Special thatnks to my lab mates, all my

friends in the Microbiology Department and to all the members of my soccer team who

made Athens the best city in the world.

Great appreciation and many thanks to my parents, who gave me unlimited love and

support throughout my schooling.

Probably the best thing that happened to me in the United States was meeting my

wife Magdalena Salameh; though research was very stressful, it was masked by her

enormous love and support. Thank you for all the kind words, support and

encouragement.

iv

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS ...............................................................................................iv

LIST OF TABLES.......................................................................................................... viii

LIST OF FIGURES..........................................................................................................ix

LIST OF ABBREVIATIONS.............................................................................................xi

CHAPTER

1 INTRODUCTION AND LITERATURE REVIEW.............................................. 1

Thermosyntropha lipolytica ......................................................................... 1

The syntrophic relationship ......................................................................... 2

What are lipases? ....................................................................................... 4

Regio-and Stereospecificity of lipases ........................................................ 5

Interface ...................................................................................................... 7

Lipase kinetics and interfacial activation ..................................................... 7

Families of bacterial lipolytic enzymes ...................................................... 10

The structure of lipases............................................................................. 15

Industrial applications of lipases................................................................ 19

Improving lipases for efficient applications................................................ 26

Lipases from extreme environment ........................................................... 28

The objectives and significance of this work ............................................. 31

References................................................................................................ 33

v

2 PURIFICATION AND CHARACTERIZATION OF TWO HIGHLY

THERMOPHILIC ALKALINE LIPASES FROM THERMOSYNTROPHA

LIPOLYTICA ............................................................................................. 83

Abstract ..................................................................................................... 84

Introduction ............................................................................................... 85

Materials and Methods.............................................................................. 86

Results ...................................................................................................... 90

Discussion................................................................................................. 93

References................................................................................................ 99

3 EFFECTS OF VARIOUS DETERGENTS ON TWO ALKALITHERMOPHILIC

LIPASES FROM THERMOSYNTROPHA LIPOLYTICA ......................... 123

Abstract ................................................................................................... 124

Introduction ............................................................................................. 125

Results and Discussion........................................................................... 126

Materials and Methods............................................................................ 132

References.............................................................................................. 137

4 OLIGOMERIZATION OF TWO THERMOPHILIC LIPASES AND ITS EFFECT

ON THERMOSTABILITY ........................................................................ 151

Abstract ................................................................................................... 152

Introduction ............................................................................................. 153

Materials and Methods............................................................................ 154

Results and Discussion........................................................................... 155

vi

References.............................................................................................. 160

5 SYNTHESIS OF FATTY ACID ESTERS AND DIACYLGLYCEROLS BY

ALKALITHERMOPHILIC LIPASES AT ELEVATED TEMPERATURES . 169

Abstract ................................................................................................... 170

Introduction ............................................................................................. 171

Materials and Methods............................................................................ 173

Results and Discussion........................................................................... 176

References.............................................................................................. 180

6 CONCLUSIONS.......................................................................................... 198

APPENDICES

A SUPPLEMENTAL FIGURES AND TABLES ............................................... 201

vii

LIST OF TABLES

Page

Table 1.1: Changes of Gibbs free energies for several hydrogen consuming and

hydrogen releasing reactions ......................................................................... 58

Table 1.2: Families of lipolytic enzymes........................................................................ 59

Table 1.3: Commercial lipases produced by Novo Nordisk ........................................... 63

Table 1.4: The most thermophilic lipases ...................................................................... 64

Table 1.5: Properties of bacterial lipases from Gram positive and Pseudomonas sp.... 65

Table 2.1: Purification of LipA and LipB produced extracellularly by Thrmosyntropha

lipolytica........................................................................................................ 107

Table 2.2: Effect of various metal ions and inhibitors on lipase activity. ...................... 108

Table 3.1: Properties of various detergents................................................................. 141

Table 3.2: The effect of E600 on LipA and LipB activity in the presence of SDS and

Tween 20...................................................................................................... 142

Table 5.1: Lipase-catalyzed esterification of fatty alcohols in various organic solvents.185

Table A.1: Metal analysis of LipA and LipB ................................................................. 202

viii

LIST OF FIGURES

Page

Figure 1.1: The lipase reaction catalyzing the hydrolysis and synthesis of lipids. ......... 68

Figure 1.2: The effect of substrate concentration on hydrolysis rate. ............................ 70

Figure 1.3: Model for describing lipase kinetics acting on insoluble substrate at the

interface.......................................................................................................... 72

Figure 1.4: The common lipase fold. ............................................................................. 74

Figure 1.5: Structure of P. aeruginosa lipase. ............................................................... 77

Figure 1.6: The secondary structure topology of BSP is compared to that of the

canonical α/β hydrolase fold ........................................................................... 79

Figure 1.7: Ribbon diagram of both lipases from G. stearothermophilus. (A) Structure of

G. stearothermophilus P1 lipase..................................................................... 81

Figure 2.1: Extracellular lipase activity ........................................................................ 109

Figure 2.2: Purified LipA and LipB proteins were analyzed by SDS–PAGE. ............... 111

Figure 2.3: Effect of temperature on activity................................................................ 113

Figure 2.4: Effect of pH on activity and stability........................................................... 115

Figure 2.5: Thermostability of LipA and LibB............................................................... 117

Figure 2.6: The activity of purified LipA and LipB toward substrates........................... 119

Figure 2.7: Positional specificity of purified LipA and LipB .......................................... 121

Figure 3.1: Temperature activity profile with 0.2% SDS .............................................. 143

Figure 3.2: The hydrolysis rate of p-nitrophenyl laurate by LipA ................................. 145

ix

Figure 3.3: Effects of various detergents on the activity .............................................. 147

Figure 3.4: The effect of detergent on stability at room temperature........................... 149

Figure 4.1: The elution profiles of aggregated LipA and LipB after gel filtration

chromatography............................................................................................ 163

Figure 4.2: 12% SDS-PAGE of eluted fractions after gel filtration of aggregated LipA

and 16% Gradient native PAGE of purified LipA and LipB ........................... 165

Figure 4.3: Effects of SDS and Tween20 on thermostability ....................................... 167

Figure 5.1: The lipase reaction catalyzing the hydrolysis and synthesis of lipids and the

synthesis of octyl oleate. .............................................................................. 186

Figure 5.2: TLC analysis of the effect of polyethylene glycol (PEG) on lipase-catalyzed

octyl oleate synthesis. .................................................................................. 188

Figure 5.3: The effect of temperature on LipA (○) and LipB (□) activity and stability in

isooctane. ..................................................................................................... 190

Figure 5.4: Time course of fatty acid alcohol esters synthesis .................................... 192

Figure 5.5: TLC analysis and time course formation of octyl oleate and lauryl oleate. 194

Figure 5.6: Time course of diacylglycerol synthesis .................................................... 196

Figure A.1: The time course of p- nitrophenyl laurate hydrolysis by LipA and LipB .... 203

Figure A.2: The catalytic activity versus enzyme concentration .................................. 205

Figure A.3: The effect of commercially available detergents on the catalytic activity of

LipA and LipB. .............................................................................................. 207

Figure A.4: SDS-PAGE of partially purified LipA and LipB. ......................................... 209

x

LIST OF ABBREVIATIONS

BCA- Bicinchoninic Acid

BSA-bovine serum albumin

CAPS- 3-(cyclohexylamino)-1-propane sulfonic acid

CMC-critical micelle concentration

CTAB- cetyltrimethyl ammonium bromide

DAG-diacylglycerols

E600- diisopropyl p-nitrophenylphosphate

EDTA-ethylenediaminetetraacetic acid

FPLC-fast protein liquid chromatography

HEPES- 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid)

HIC-hydrophobic interaction chromatography

IEC-ion exchange chromatography

LCFA-long chain fatty acids

LipA-lipase A

LipB-lipase B

NEFA-non esterified fatty acids

NPN- N-phenyl-1-naphthylamine

PAGE-polyacrylamide gel electrophoresis

PEG-polyethylene glycol

PMSF-phenylmethanesulfonoylfluoride

xi

pNP-para nitrophenol

pNPL-para nitrophenyl laurate

pNPP-para nitrophenyl palmatate

pNPB-para nitrophenyl butyrate

TAG-triacylglycerol

TAPS- n-tris(hydroxymethyl)methyl-3-aminopropanesulfonic Acid

TLC-thin layer chromatography

Tris- tris [hydroxymethyl] aminomethane

Triton X-100- t-octylphenoxypolyethoxyethanol

Tween 20- polyoxyethylene sorbitan monolaurate

Tween 80- polyoxyethylene sorbitan monooleate

xii

CHAPTER 1

INTRODUCTION AND LITERATURE REVIEW

Thermosyntropha lipolytica

The lipases described in this thesis are produced by Thermosyntropha lipolytica. It is

an anaerobic thermophilic organotrophic lipolytic alkalitolerant bacterium. It was isolated

with a specific intent of finding bacterial lipases with tolerance for, and activity, at high

temperatures and high pH values. This bacterium was isolated from alkaline hot springs

of Lake Bogoria, Kenya, on minimal media plus olive oil. The optimum pH (measured at

25 °C) is between 8.1 and 8.9 and temperature optimum is between 60 and 66 °C

determined at optimum pH (Svetlitshnyi, et al., 1996).

The cells stain Gram negative but the organism is a Gram-type positive (Wiegel,

1981) bacterium. This is consistent with the 16S rRNA sequence analysis, which placed

the organism into the Clostridium-Bacillus subphylum (Firmicutes) and identified

Syntrophospora and Syntrophomonas spp. as closest phylogenetic neighbors.

In contrast to most lipolytic microorganisms T. lipolytica does not utilize the liberated

glycerol. Although the lipase activity is constitutive, T. lipolytica was able to grow on

long-chain fatty acids only in syntrophic co-culture with Methanobacterium, it grew on

triacylglycerols and linear saturated and unsaturated fatty acids with 4 to 18 carbon

atoms. In the absence of a methanogen, the addition of olive oil, soybean oil or any

1

triacylglycerol did not stimulate the growth on basal medium containing yeast extract.

The triacyl glycerides were hydrolyzed by a constitutively formed lipase found in the

culture supernatant, but the liberated glycerol and long-chain fatty acids were not

significantly utilized regardless of whether CaCl2 was added in equimolar concentrations

relative to the fatty acids content of the triacylglycerides because the degradation is

energetic unfavorable until the formed H2 is removed to nM concentrations rendering

the reaction exergonic (Svetlitshnyi, et al., 1996).

Several anaerobic bacteria which are able to utilize fatty acids in syntrophic

relationship have been isolated and characterized such as: Syntrophomonas wolfei

subsp. wolfei DSM 2245T (McInerney, et al., 1979, 1981), Syntrophospora bryantii DSM

3014T (Stieb and Schink, 1985), Syntrophomonas sapovorans DSM 3441T (Roy, et al.,

1986), Syntrophomonas wolfei subsp. saponavida DSM 4212T (Lorowitz, et al., 1989).

Syntrophomonas curvata DSM 15682T (Zhang, et al., 2004). Syntrophothermus

lipocalidus DSM 12680T is the only other syntrophic thermophilic bacterium recently

isolated, however it is not described as a lipase producer, since the strain did not utilize

any triglycerides (Sekiguchi et al., 2000). Until now, no other thermophilic anaerobe

responsible for the degradation of triglycerides had been identified.

The syntrophic relationship

Syntrophism is a special symbiotic relationship between two metabolically different

microorganisms which depend on each other for the degradation of certain substrates.

Methanogenic bacteria can degrade primarily only one-carbon compounds. Therefore,

acetate, propionate, ethanol, and their higher homologs have to be fermented further to

2

one-carbon compounds. These fermentations are called secondary or syntrophic

fermentations.

It was shown that ethanol fermentation by Methanobacillus omelianki is in fact

carried out by a methanogen and a non methanogenic species in a syntrophic

association (Bryant, et al., 1967, 1977).

Ethanol is fermented by bacteria according to Eq. 1

2CH3CH2OH + 2H2O ↔ 2 CH3COO- + 4 H2 + 2 H+ ………. (1)

∆G°′ = 19.2 kJ/ reaction

This fermentation is thermodynamically unfavorable unless the pressure of hydrogen

gas is decreased to a lower level by a methanogen

4 H2 + HCO-3 + H+ ↔ CH4 + 3H2O ………. (2)

∆G°′ = -135.6 kJ/ reaction

It was also suggested that fatty acids with more than two carbons are degraded by

methanogens and another group of bacteria like the ethanol degrader and produce

acetate and hydrogen. Later, many anaerobic bacteria that degrade butyrate and longer

fatty acids were isolated and characterized in co-culture with methanogens (Bryant, et

al., 1967; Jackson, et. al., 1999; McInerney, et al., 1981; McInerney, et al., 1979). The

degradation of fatty acids to acetate, hydrogen and CO2 is eight times more endergonic

than the ethanol oxidation and subsequently, the hydrogen pressure should be

decreased at a much lower level (<10 Pa) than ethanol (<100Pa) (Table1.1) (Schink, et

al., 1997).

3

What are lipases?

Lipases (E.C. 3.1.1.3) are carboxyl ester hydrolases that catalyze the hydrolysis and

synthesis of long chain acylglycerols (Fig. 1.1) (Sarda, et al., 1958; Ferrato, et al., 1997).

There are interesting features that distinguish lipases from other hydrolases and

especially esterases. Lipases are activated at the water-lipid interface, they show little

activity when the substrate is in the monomeric form and their activity increases

dramatically above the solubility limit where lipids start to form emulsions. This fact has

led to the emergence of a phenomenon known as interfacial activation, which describes

emulsions as a necessity for the lipolytic reaction to occur (Sarda, et al., 1958; Ferrato,

et al., 1997).

For most enzymes, their active sites have easy access to the solvent; this is not

exactly the case for most lipases, where their active sites are covered by a hydrophobic

lid that greatly constricts the accessibility of substrates. The lid is basically a surface

loop covering the active site of the enzyme and moves away on contact with the water-

lipid interface (Brzozowski, et al., 1991; Derewenda, et al., 1992). Several

crystallography studies have suggested that a conformational change has to occur so

that substrate is able to access the active site (Brady, et al., 1990; Winkler, et al., 1990).

The elucidation of many other 3D structures of lipases complexed with substrates and

inhibitors have provided an elegant explanation of interfacial activation; at the absence

of substrate, the active site of the lipase is covered by the lid which creates the closed

(inactive) conformation. This is the dominant conformation in aqueous media and in the

absence of the interface with as indicated by a low lipolytic activity of lipases. Once the

substrate is available, the enzyme undergoes substantial conformational changes. The

4

enzyme binds to the interface triggering the lid to move away turning the closed form of

the enzyme into the open active form where the active site is exposed to the substrate

and accessible to the solvent (Brzozowski, et al., 1991). The movement of the lid not

only opened access to the active site, but also buries a large portion of the hydrophilic

area in association with the exposure of larger hydrophobic parts to the solvent

(Brzozowski, et al., 1991; Derewenda, et al., 1992; van Tilbeurgh, et al., 1993).

The European BRIDGE-T-Lipase project has revealed few 3D structures and

numerous biochemical and kinetic data. It was concluded from these studies that not all

lipases show interfacial activation. Some of them, such as the lipases from

Pseudomonas aeruginosa and Burkholderia glumae (bas Pseudomonas glumae), have

an amphiphilic lid covering their active sites (Verger, 1997), while others, such as the

lipases from Bacillus subtilis 168 (Lesuisse, et al., 1993), Fusarium solani (cutinase with

lipolytic activity) (Hjorth, et al., 1993) and guinea pig pancreas (Martinez, et al., 1992)

have no lid.

The discoveries of these exceptional lipases led to the conclusion that the presence

of a lid domain and interfacial activation are unsuitable criteria to classify an enzyme as

a lipase. Therefore a lipase is simply a carboxylesterase that catalyses the hydrolysis of

long chain acylglycerols (Verger, 1997).

Regio-and Stereospecificity of lipases

Lipases have been found to exhibit varying degrees of specificity for the positioning

of the ester bond on the alcohol backbone, referred to as regioselectivity. Although the

glycerol molecule has plane symmetry, the two primary alcohol groups are sterically

distinct, and substitution of these hydroxyl groups with different acyl groups leads to

5

optically active compounds. The glycerol molecule is conventionally written in the Fisher

projection with the secondary alcohol group to the left and the two primary alcohol

groups to the right, with the backbone carbons numbered 1, 2, and 3 from top to bottom.

Many microbial lipases preferentially cleave the ester bonds at the primary alcohol

positions (sn-1 and sn-3, for stereospecifically numbered glycerol). Furthermore, there

is usually specificity for the chain length of fatty acid and the degree of saturation, and

thus the regioselectivity of the enzyme may be more or less pronounced depending on

fatty acid chain length (Jaeger, et al., 1994; Kazlauskas, 1994). The regiospecific

lipases offer the greatest potential for industrial application, such as production of

structured lipids with unique functional properties (Sonnet and Gazzillo, 1991). The

stereo-enantioseletivity enables the enzyme to discriminate between the enantiomers of

a racemic pair. This property is very important in the synthesis of fine chemicals,

because of its ability to produce at least a highly enriched active enantiomer instead of a

mixed racemic compound.

Most lipases can act on several different kinds of acyl-alcohols, which include

specificity as well for the alcohol molecule. The reverse, i. e., synthesis reaction is

energetically favored in water-restricted environments with enzyme specificities and

preferences largely retained. Thus, the synthesis of specific acyl-alcohol molecules can

be envisioned by setting the correct conditions in the aquous solution (see below).

Enantiomeric specificity, especially in transesterification reactions, allows production of

specific chiral esters, which are important intermediates in production of

pharmaceuticals and pesticides, as well as for the optical purification of racemic

mixtures.

6

Interface

Triacylglycerols (TAG) are uncharged lipids. Although those with short chain fatty

acids are slightly soluble in water, compounds with longer chain fatty acids are insoluble.

The maximum concentration of monomers in aqueous solution is called the saturation

value. This is the point where triacylglycerols start to form emulsions. Phospholipids and

most surfactants are amphipathic compounds. It means that they contain both a

hydrophobic “tail” and a hydrophilic “head group”. When placed in an aqueous

environment, amphipathic molecules aggregate in such a way as to expose the polar

head group toward water and will therefore be attracted to the surface of the polar

phase. They form micelles when exceeding the maximum concentration of dissolved

monomer at a point called the critical micelle concentration (cmc) (for quantitative

measurements see chapter 3, table 3.1) (Jaeger, et al., 1994). The enzymatic

catalyzed lipolysis occurs exclusively at the lipid-water interface, implying that the

concentration of substrate molecules at the interface is expressed in mol m-2 and

directly determines the rate of lipolysis.

Lipase kinetics and interfacial activation

The challenge of studying lipase kinetics is that their common substrates

(triglycerides) are insoluble in aqueous solution even at µM concentrations. The

interfacial km, in which the concentration is expressed as moles per unit area instead

per volume, is difficult to interpret. It may be more an estimate of the relatively non

specific protein-lipid interaction than an actual measure of substrate specificity (Woolley

and Peterson, 1994). Thus, despite being a single substrate enzyme reaction, one

cannot expect Michaelis-Menten kinetics, and this is a fundamental difference between

7

esterases and lipases. Esterases exhibit Michaelis-Menten kinetics and react with

soluble substrates (triacylglycerols with short chain fatty acids; e.g. triacetin). They have

low activities toward insoluble glycerides (triacylglycerols with long chain fatty acids; e.g.

trilaurin). The maximum reaction rate is reached well before the saturation point (Figure

1.2). The lipases, in contrast, are active toward insoluble substrates, and have generally

a negligible activity toward soluble short chain fatty acid esters (Sarda, et al., 1958;

Brzozowski, et al., 1991; Derewenda, et al., 1992 and Jaeger, et al., 1994). Although

there are a few exceptions where lipases show as high as 90% relative activity toward

substrates in their monomeric form compared to emulsions and micelles (Nini, et al.,

2001).

Chahinian and coworkers (2002) compared the kinetic behavior of carboxyl ester

hydrolases against solutions and emulsions of vinyl esters and triacylglycerols (TAG) to

allow better distinction between lipases and esterases. They showed that esterases

display maximal activity against solutions of short-chain vinyl esters (vinyl acetate, vinyl

propionate, and vinyl butyrate) and short chain TAG (triacetin, tripropionin, and

tributyrin). Half-maximal activity is reached at ester concentrations far below the

solubility limit. The transition from solution to emulsion at substrate concentrations

exceeding the solubility limit has no effect on esterase activity. Lipases are also active

on solutions of short-chain vinyl esters and short chain TAG. However, in contrast to

esterases, they all display maximal activity against emulsified substrates and half-

maximal activity is reached at substrate concentrations near the solubility limit of the

esters. The kinetics of hydrolysis of soluble substrates by lipases are also hyperbolic

and show no or weak interfacial activation (Chahinian, et al., 2002).

8

Verger and coworkers (1973) proposed a kinetic model for the action of lipases

at interfaces. The model proposes a reversible adsorption or penetration process of the

lipases to the interface, and this may include the activation of the enzyme (moving the

lid away and exposes the active site). This is regarded as the rate limiting step in

contrast to the ternary/quaternary complex of enzymatic reaction with soluble substrates.

The adsorption is followed by the formation of the enzyme-substrate complex which

leads them to the catalytic step of hydrolyzing the ester and forming the product and the

enzyme for further catalysis as in Michaelis-Menten fashion (figure 1.3A).

Because of the diversity of applications for lipases, their kinetics have been

extensively studied. Various kinetic mechanisms were proposed to describe lipase-

mediated synthetic reactions; for example, the Michaelis-Menten uni-uni mechanism

and ping-pong bi bi mechanism (Figure 1.3B) (Paiva, et al., 2000; Malcata, et al., 2000).

A ping-pong bi-bi mechanism was also proposed to characterize the kinetics of glucose

acylation with fatty acids in acetone (Arcos, et al., 2001) and 2-ethyl-2-butanol (Flores,

et al., 2002).

The following general chemical formula, originally adopted by Deems et al. (1975)

for phospholipase, can be used in case where detergents are used, and micelles are

involved in binding to and activating the enzyme assuming that all micelles have the

same strong affinity to the enzyme, and no inhibition exist other than substrate surface

area constrains:

9

Where [A] and [B] can be calculated as following (Abousalham, et al., 1997)

x[P] + y[D]

[A] = n

x[P] [B] =

x[P] + y[D]

where A is concentration of enzyme adsorption sites, B is substrate surface fraction, v is

the initial velocity (U·mg-1), Vmax is the maximum velocity (U·mg-1), x is the average

surface area of substrate (cm2/mol), y is the average surface area of detergent

(cm2/mol), [P] is substrate concentration (M), [D] is detergent concentration (M), n is the

micellar surface area (cm2/mol),

KSA = , Km

B = k-2 + k3 k-1

k2

Families of bacterial lipolyt

Although lipolytic enzyme

high sequence homology an

molecular mass which can

marcescens). For systematic

function of this rapidly grow

Database (LED) has been d

information on sequence, s

proteins and then assigned to

similarity (http://www.led.uni-s

k1

ic enzymes

s in general have very conserved 3-D structures but lack a

d display a wide diversity of properties including their

range from 19 KDa (LipA of B. subtilis) to 65 KDa (S.

analysis of the relationship of sequence, structure, and

ing, highly diverse protein class, the Lipase Engineering

esigned to serve as a navigation system that integrates

tructure, and function of lipases, esterases, and related

homologous families and subfamilies based on sequence

tuttgart.de) (Fischer and Pleiss, 2003).

10

Newly discovered bacterial lipases were classified into any of the three

pseudomonas groups, based mainly on their size and desirable industrial properties,

such as the regiospecificity of hydrolyzing triglycerides (Jaeger, et al., 1994). However,

this classification system was too primitive and ineffective, because of the wide diversity

of many new lipases being constantly identified from different genera. In 1999, a new

classification of bacterial lipases and esterases was published. Based on comparison

of their amino acids sequences and some fundamental biological properties, bacterial

lipases were classified into eight families (Arpigny and Jaeger, 1999) discussed below:.

Family I. The true lipases

This family was originally classified into six subfamilies by (Arpigny and Jaeger,

1999), based on physical properties and amino acids sequences. However, the authors

of this review believe that Staphylococcal lipases should not be included in the same

subfamily as of thermophilic lipases from Geobaillus sp. because of low amino acid

similarities (28%) and vast differences in physical and molecular properties. Therefore,

the authors propose to introduce a seventh subfamily that only contains lipases from

Staphylococcus sp.

Subfamily I.1. contained originally P. aeruginosa lipase and then lipases from

Vibrio cholerae, Acinetobacter calcoaceticus, P. wisconsinensis and Proteus vulgaris

were included which have molecular masses in the range of 30–32 KDa and display a

high sequence similarity to the P. aeruginosa lipase.

Subfamily I.2. contains lipases from Burkholderia cepacia, Burkholderia glumae,

and Chromobacterium viscosum share several structural features. They are

characterized by a slightly larger size (33 kDa) than subfamily I.1 because of an

11

insertion in the amino acid sequence forming an anti-parallel double β-strand at the

surface of the molecule (Noble, et al., 1993). One important feature that both I.1 and I.2

subfamilies share is that the expression of active lipase depends on a chaperone

protein named lipase-specific foldase (‘Lif’). In addition, a Ca2+-binding site and a

disulphide bridge are conserved in a majority of sequences.

Subfamily I.3. contains lipases from at least two distinct species: P. fluorescens

and Serratia marcescens. They have in common a higher molecular mass than lipases

from subfamilies I.1 and I.2 (Ps. fluorescens, 50 kDa; S. marcescens, 65 kDa) and the

absence of N-terminal signal peptide. The secretion of these enzymes occurs in one

step through a three-component ATP-binding-cassette transporter system (Li, et al.,

1995; Duong, et al., 1994)

Subfamily I.4. contains lipases from Gram type positive Bacillus species. The

feature that Bacillus and Geobacillus (bas Bacillus) species have in common is that an

alanine residue replaces the first glycine in the conserved pentapeptide: Ala-X-Ser-X-

Gly, an exceptional enzyme that lacks the conserved pentapeptide is LipA of B. subtilis

168 (Dartois, et al., 1992).

Many of the discovered mesophilic Bacillus lipases belong to this subfamily including

LipA (Li, et al., 1995) and LipB (Eggert, et al., 2000) of B. subtilis, B. pumilus (Kim, et al.,

2002), and Bacillus sp. Bp-6 (Ruiz, et al., 2003) and Bacillus megaterium (Ruiz, et al.,

2002). They are the smallest lipases described so far (around 20 kDa) and show high

homology among each other (B. subtilis lipases have 75% and 98% identity to B.

pumilus and B. megaterium, respectively).

12

Subfamily I.5. contains thermophilic lipases with similar properties from

Geobacillus thermocatenulatus, Geobacillus stearothermophilus and Geobacillus

thermoleovorans. Their maximal activity is at pH 9.0 and 65-75 °C. Their molecular

mass is approx. 45 kDa (Rua Luisa , et al., 1997; Schmidt-Dannert, et al., 1994; Kim, et

al., 1994; Sinchaikul, et al., 2001; Cho, et al. 2000).

Subfamily I.6. contains lipases from the major bacterial inhabitants of human

skin. Propionibacterium acnes (33 kDa) (Miskin, et al., 1997) and Streptomyces

cinnamoneus (29 KDa) (Sommer, et al., 1997). They show 50% similarity to each other

including the central region (normally contain the conserved pentapeptide) which is

approx. 50% similar to lipases from B. subtilis and they share only 15% similarity to

subfamily I.1.

Subfamily I.7. contains nine lipases from six Staphylococcus species, three from

S. epidermidis, two from S. aureus, and one each from S. haemolyticus, S. hyicus, S.

warneri, and S. xylosus, have been determined (Rosenstein, et al., 2000). All are

similarly produced as pre-pro-proteins, with pre-regions corresponding to a signal

peptide of 35 to 38 amino acids, a pro-peptide of 207 to 321 amino acids, and a mature

peptide comprising 383 to 396 amino acids (Rosenstein, et al., 2000; Gotz, et al. 1998).

They all show high similarity ranging from 64% to 88%. The lipases are extra-cellularly

secreted in the pro-form and are afterwards cleaved to the mature form by specific

proteases. The propeptide presumably acts as an intramolecular chaperone which

facilitates the translocation of the lipase across the cell membrane. It was also observed

that the pro-region protects the proteins from proteolytic degradation. All staphylococcal

lipases are Ca2+ dependent. However, despite being very similar in their primary

13

structures the staphylococcal lipases show significant differences in their biochemical

and catalytic properties, such as substrate selectivity, pH optimum and interfacial

activation. Moreover, the lipase from S. hyicus is unique by displaying high phospho-

lipase activity (Rosenstein, et al., 2000; Gotz, et al. 1998).

Family II. GDSL

This family contains lipases that do not exhibit the conventional pentapeptide

Gly-X-Ser-X-Gly but rather display a Gly-Asp-Ser-(Leu) (GDSL) motif. Enzymes belong

to this family are mostly esterases, such as the esterases from Aeromonas hydrophila,

P. aeruginosa, Salmonella typhimurium and Photorhabdus luminescens (Arpigny and

Jaeger, 1999).

Family III

Members of this family are the extracellular lipases of Streptomyces exfoliatus and

Streptomyces albus. S. exfoliates lipase show 20% identity to two mammalian platelet-

activating factor acetylhydrolases (PAF-AHs) (Wei, et al., 1998).

Family IV HSL

This family contains lipases and esterases with significant sequence similarity to

mammalian hormone-sensitive lipase (HSL). Hormone-sensitive lipase is the key

enzyme in the mobilization of fatty acids from adipose tissue, sequence alignments

have failed to detect any significant homology between hormone-sensitive lipase and

the rest of mammalian lipases and esterases. However, a remarkable secondary

structure homology between HSL and bacterial and fungal lipases were found

(Contreras, et al., 1996). Some members of this family are esterases from psychrophilic

14

Moraxella sp., Psychrobacter immobilis, mesophilic Escherichia coli, Alcaligenes

eutrophus and thermophilic Alicyclobacillus acidocaldarius, Archeoglobus fulgidus.

Families V, VI, VII and VIII

Members of these families are rare. The families mainly contain esterases (see Table

1.2.).

The structure of lipases

An inceasing number of bacterial and fungal 3-D structures of lipases and esterases

have been solved in recent years. The first 3-D structure of a lipases started was

reported in 1979 by Hata and coworkers who obtained a low resolution structure of the

enzyme from the fungus Geotrichum candidum (Hata, et al., 1979). It was not until 1990

that the first high resolution structure of a fungal lipase was solved, that from

Rhizomucor miehei (Brady, et al., 1990), and this was followed by structures of the

enzyme from human pancreas (Winkler, et al. 1990) and higher resolution Geotrichum

candidum (Schrag, et al., 1991). The first crystal structure elucidated of bacterial lipase

was the one from Burkholderia (bas Pseudomonas) glumae (Noble, et al., 1993). In

recent years, many crystal structures of lipases from bacterial origin were elucidated

(Lang, et al., 1996; Kim, et al., 1997; van Pouderoyen, et al., 2001; Jeong, et al., 2002),

including lipase structures from the alkalithermophilic bacteria Geobacillus (bas Bacillus)

stearothermophilus strains L1 (Jeong, et al., 2002) and P1 (Tyndall, et al., 2002).

The elucidation of these crystal structures of lipases has dramatically increased our

knowledge of their catalytic mechanism. All the lipases investigated so far vary

considerably in size and in their amino acid sequences. Despite the remarkable tertiary

structural homolog from widely different systems, i.e. presence of the hydrophobic lid

15

and the triads of amino acids in the catalytic center, there is little sequence similarity

among lipases in general (Bell, et al., 1999).

The studies of the 3-D structures of lipases have shown interesting findings. The

active site was found to consist of a Ser-His-Asp/Glu catalytic triad reminiscent of the

serine proteases (Cygler and Schrag, 1997), and an unusual feature of this active site is

that it is not exposed on the protein surface, instead, it is completely buried under a lid

like structure composed of α-helices, and is not accessible to the substrate. This led to

the hypothesis that lipases undergo a significant conformational change once adsorbed

to the lipid-water interface allowing the substrate to be able to access the active site.

Another significant conformational change involves the increase of the hydrophobic

surface of the enzyme, which is involved in the lipid recognition (Jaeger, et al., 1994).

The 3-D structures also revealed basic fold pattern of many lipases. They belongs to the

α/β hydrolase fold family. This fold of enzymes is one of the largest groups of

structurally related enzymes with diverse catalytic functions. Beside lipases, members in

this family include esterases, cholinesterases, cutinases, haloalkane dehalogenase,

endopeptidase, serine carboxypeptidase, proline aminopeptidase, proline

oligopeptidase, haloperoxidase hydroxynitrile lyase, and epoxide hydrolase (Wei, et al.,

1999; Bourne, et al., 2004; Nagy, et al., 2003; Holmquist, et al., 2000; Nardini and

Dijkstra, 1999; Schrag and Cygler, 1997; Ollis, et al., 1992).

Many lipase structures have been solved, although these proteins show little

sequence homology, all are members of this fold family (Ollis, et al., 1992). Lipases

span a wide range of molecular weights, from 19 kDa to 60 kDa. Their secondary

structure is generally composed of several (up to eight) parallel β sheets (β1-β8)

16

connected by α helices (up to six) (Figure 1.4A) (Ollis, et al., 1992; Schrag and Cygler,

1997).

The active site is composed of side chains of three amino acids serine, aspartic or

glutamic acid and histidine, which form the generally conserved catalytic triad for the

nucleophilic attack. Catalysis starts by the serine oxygen on the carbonyl carbon atom

of the ester bond, forming the tetrahedral intermediate stabilized by hydrogen bonding

to the nitrogen atoms of main chain residues. This leads to the formation of an acyl-

lipase complex, which leads to the release of the product, the free fatty acids, and

generation of the free enzyme (Figure 1.4B). The formation of the tetrahedral

intermediate and its stabilization by hydrogen bonding was first postulated for serine

proteases (Brady, et al., 1990; Winkler, et al. 1990; Blow, et al., 1969) and known as

the oxyanion hole (Figure 1.4B). The nucleophilic Ser residue is normally found in a

highly conserved penta peptide G-X-S-X-G (X represent any amino acid) located at the

C-terminal end of strand β5, forming a β-turn-α motif named the ‘nucleophilic elbow’

(Figure 1.4A). In lipases from Geobacillus and Bacillus species, the first glycine in the

conserved penta peptide is an alanine. As discussed above most active sites of the

bacterial lipases are covered by a hydrophobic lid that maintains the catalytic triad in a

hydrophobic environment inaccessible to the solvent. The topological location,

complexity and length of the lid vary among the lipases (Schrag and Cygler, 1997).

To date, eight bacterial true lipase 3-D structures have been solved; Burkholderia

(bas Pseudomonas) glumae (Noble, et al., 1993), Chromobacterium viscosum (Lang,

et al., 1996), Burkholderia (bas Pseudomonas) cepacia (Kim, et al., 1997; Schrag, et al.,

1997), Pseudomonas aeruginosa (Nardini, et al., 2000), Bacillus subtilis (van

17

Pouderoyen, et al., 2001), Streptomyces exfoliatus (Wei, et al., 1998), and the first two

alkalithermophilic lipases from Geobacillus (bas Bacillus) stearothermophilus L1 (BSL)

(Jeong, et al., 2002) and Geobacillus stearothermophilus P1 (BSP) (Tyndall, et al.,

2002). Other solved structures are from lipase related bacterial enzymes such as;

Streptomyces scabies esterase (SsEST) (Wei, et al., 1995), Alcaligenes eutrophus

esterase (Bourne, et al., 2000) and Pseudomonas flurorescens carboxylesterase (Kim,

et al., 1997).

The first bacterial structure solved as mentioned earlier was the lipase of

Burkholderia glumae. The enzyme contains three domains, the largest of which

contains a subset of the α/β hydrolase fold and a calcium site. It also contains the lid

(found on α5) which controls substrate access to the active site. The lipase of B. cepacia

shares several structural features with homologous lipases from B. glumae and

Chromobacterium viscosum, including a calcium-binding site. In contrast, the C.

viscosum lipase contains an oxyanion hole similar to serine proteases (Noble, et al.,

1993).

The structure P. aeruginosa reveals a highly open conformation with a solvent-

accessible active site. This is in contrast to the structures of B. glumae lipase and C.

viscosum lipase in which the active site is buried under a closed or partially opened lid

(Lang, et al., 1996).The lipase structures from B. glumae, B. cepacia, and C. viscosum,

show 42% amino acid sequence identity to P. aeruginosa lipase. The structural similarity

is mainly localized in the core domain. The α helix 5 and its neighboring loops form the

lid. Compared with the canonical α/β hydrolase fold, the first two -strands are absent,

and therefore, to be consistent with the numbering of the consensus α/β hydrolase fold,

18

the first strand in the P. aeruginosa structure is named 3 (Figure.1.5) (Nardini, et al.,

2000).

The two lipases from Geobacillus stearothermophilus (BSP) and (BSL) provided the

first structures of lipases from a thermophilic microorganism. BSL show high sequence

identity with BSP ( 95%). All possess a complete sequence of 417 residues including a

29-residue signal sequence that is cleaved to produce the mature 388-residue lipase.

The optimum activity of both thermostable lipases lies around 65 °C and pH 8.0–9.0.

They share less than 20% amino acid sequence identity with other lipases for which

there are crystal structures, and both enzymes show significant structural differences

when compared with the typical α/β hydrolase canonical fold because of numerous

insertions and deletions throughout the structures (Figure. 1.6). Both structures contain

zinc and calcium binding sites which is unique among all lipases. Zinc binding is

mediated by two histidine and two aspartic acid residues and may play a role in thermal

stability. The catalytic triad is identical for both enzymes and covered by a lid like

structure (long helix α6) (Figure 1.7) (Jeong, et al., 2002; Tyndall, et al., 2002).

Industrial applications of lipases

Lipases from bacterial and fungal origin are the most widely used in various

biotechnological applications. In general, they are stable in wide range of organic

solvents and many show high thermostability, they have wider pH and temperature

optima range than lipases from eukaryotic origin, they do not require cofactors and they

have a very diverse substrate range with high regio- and enantioselectivity making

microbial lipases an important and attractive choice for many applications in organic

synthesis. The large biotechnological versatility of lipases is based on their ability to

19

catalyze the hydrolysis and various reverse reactions such as esterification,

transesterification, and aminolysis in organic solvents. They are involved in industries

like pharmaceutical, dairy, detergent, cosmetic, oleochemical, fat processing, leather,

textile, cosmetic, paper industries and others. Below follows an overview of the

applications of lipases and their biotechnological importance (Jaeger, et al., 1994;

Bornscheuer, et al., 2002; Schmidt-Dannert, 1999; Jaeger, et al., 1999; Villeneuve, et

al., 2000; Balcão, et al., 1996; Jaeger and Reetz, 1998).

Medical biotechnology

Lipases have been used in the medical field, particularly in the treatment and/or

development of atherosclerosis and hyperlipidemia, and its importance in regulation and

metabolism, since products of lipolysis such as free fatty acids and diacylglycerols play

many critical roles, especially as mediators in cell activation and signal transduction

(Farooqui, et al., 1987). Lysosomal lipases show optimal activity at an acidic pH (around

pH 5.0), whereas plasma membrane and microsomal lipases are optimally active at pH

7.0. A deficiency of acid lipase results in Wolman's and cholesterol ester storage

diseases. Both of these diseases are characterized by intralysosomal accumulation of

triacylglycerols and cholesterol esters. Patients are diagnosed by determining acid

lipase activity in their leukocytes and fibroblasts. Moreover, lipases were found to have

a strong relation with the development of tumors (Quigley, et al., 2001; Mamputu and

Renier, 1999).

Lipases are increasingly on demand for nutraceuticals industry. A nutraceutical is

any substance that can be considered a food that provides medical or health benefits,

including the prevention and/or treatment of a disease. For instance, a lipase from

20

Pseudomonas sp. immobilized within the walls of a hollow-fiber reactor was used to

convert the linoleic acid present in corn oil into conjugated linoleic acid (CLA) which is a

nutraceutical that has anti-carcinogenic and anti-atherogenic activities (Sehanputri, et

al., 2000).

Obesity is regarded as one of the top health issues in the world; it is a serious

medical disease that affects over a quarter of adults in the United States, and about

14% of children and adolescents according to America Obesity Association (AOA)

(http://www.obesity.org/). One of the possible treatments for obesity is developing lipase

inhibitors. Orlistat is the first agent in the lipase inhibitor class of anti-obesity drugs; it

has been shown to reduce body weight by inhibiting absorption (by approximately 30%)

of ingested dietary fat and was proposed as a novel approach for obesity treatment

(Lucas, et al., 2001).

Indomethacin, ketoprofen and etodolac are non-steroidal anti-inflammatory drugs

(NSAIDs). Their therapeutic efficacy is often limited because of their poor aqueous

solubility and permeability (Akimoto, et al., 2003). To increase their solubility, various

methods have been used, including salt formation, dispersion with a polymer and the

pro-drug approach. Among all these methods the pro-drug approach seems very

promising. Forming esters with sugar using lipases is effective in enhancing drug

solubility, and is quite effective in preparing pro-drugs. Wang and co workers used a

lipase from Candida antarctica to catalyze the transesterification of glucose with vinyl

esters of indomethacin, ketoprofen and etodolac (Wang, et al., 2005). Trans-

esterification (acidolysis) was successfully done in n-hexane by several microbial

lipases.

21

Detergent industry

An estimated 1000 tons of lipases are added to approximately 13 billion tons of

detergents produced each year (Godfry and West, 1996). The inclusion of lipase in

detergents, one of the most important applications of lipases, includes household and

industrial laundry detergents and household dish-washers detergents. There is no

recent estimate of the total world lipase sale or the sales of detergent enzymes.

However, enzyme sales in 1995 have been estimated to be US$ 30 million of which US

$10 million was for detergent enzymes (Jaeger and Reetz, 1998).

Novo Nordisk, the leading company for industrial enzymes, introduced many lipases

as detergent additives among other various industrial applications Table1.3. They

produced the first commercial lipase named LipolaseTM from T. lanuginosus and it was

used in the detergent industry. Other bacterial lipases like LumfastTM from P. mendocina

and LipomaxTM from P. alcaligenes were produced by Genencor International and used

also as detergent additives (Jaeger, et al., 1999).

Organic synthesis

Lipases in organic synthesis are primarily used to catalyze enantioselective

reactions for the synthesis of fine chemicals and especially in preparing chiral

intermediates for pharmaceuticals (Patel, 2000). It is well recognized that therapeutics,

agrochemicals and flavor compounds are quite difficult to synthesize with chemical

methods, especially when only one enantiomer drug out of two is functional (Jaeger and

Eggert, 2002).

In the field of therapeutics, lipases are used in the synthesis process of epothilones,

which are macrolide natural products exhibiting potent anti-tumor activity against a wide

22

spectrum of human tumor cell lines including multi-drug resistant cell lines (Broadrup, et

al., 2005). A highly enantioselective lipase from Pseudomonas species was used to

catalyze the introduction of chirality at C15 position in the synthesis of epothilone D

(Broadrup, et al., 2005) and epothilone A (Zhu and Panek, 2000, 2001).

Candida rugosa lipase catalyzes the enzymatic resolution of the antimicrobial

compounds (S)- and (R)-elvirol and their derivatives (S)-(+)- and (R)-(−)-curcuphenol

(Ono, et al., 2001). The lipase Novozym 435 from Candida antarctica B was used to

catalyze acylation of the immunosuppressant and antifungal agent rapamycin and 42-

ester derivatives (e.g. rapamycin 42-hemisuccinate, 42-hemiadipate) with various

acylating agents with complete regio-selectivity and high yields (Gu, et al., 2005). The

same acylation approach was also conducted to synthesize temsirolimus (CCI-779), a

compound currently under development as a tumor inhibitor (Shaw, et al., 2001). The

same enzyme was used in the synthesis of Lubeluzole, a drug for the acute treatment of

ischemic stroke (Liu, et al., 2001).

Biodiesel production

Biodiesel, a monoalkyl fatty acid ester (preferentially methyl and ethyl esters) is

presently evaluated as a replacement for diesel. Biodiesel has a few advantages over

petroleum diesel; it is biodegradable; its combustion products have reduced levels of

particulates, carbon oxides and sulfur oxides (Iso, et al., 2001). The conversion of

vegetable oil to methyl- or other short-chain alcohol fatty acids esters can be catalyzed

in a single transesterification reaction using lipases in organic solvents (Eq. 1).

23

CH2-OOC-R1 R1-COO-R′ CH2OH

CH-OOC-R2 + 3R′ R2-COO-R′ + CH2OH …….. (1) Lipase

CH2-OOC-R3 R3-COO-R′ CH2OH

Triacylgycerol Alcohol Fatty acids esters Glycerol

Lipases tested for the production of biodiesel include the enzymes from Rhizomucor

miehei (immobilized on ion-exchange resins) and Thermomyces lanuginosa

(immobilized on silica gel). They were used to convert sunflower oil into biodiesel by

methanolysis (Al-Zuhair, 2005). Lipases from Pseudomonas cepacia (Noureddini, et al.,

2005) and Novozym 435 (Du, et al., 2004) were used to convert soybean oil. Several

processes for biodiesel fuel production have been developed, among which

transesterification using alkali-catalysis gives high levels of conversion of triglycerides to

methyl esters in short reaction times. This process has therefore been used for

biodiesel fuel production in a number of countries, including Belgium, France, Germany,

Italy and United states, with fuel production exceeds 100,000 tons yearly (Fukuda, et al.,

2001). The major disadvantage of biodiesel production is the coast of biocatalysts

(Jaeger and Eggert. 2002). One possible solution to reduce cost is to enhance their

stability and use immobilization technology to reduce coast (Iso, et al., 2001).

Agrochemical industry

In the agrochemical industry, lipases have been used in the synthesis of herbicides.

Indanofan is a novel herbicide used for grass weeds in paddy fields. It was

commercialized as a racemic mixture in 1999; however, by examining the herbicidal

activity of each enantiomer, only the (S)-enantiomer is active. To synthesize this

enantiomer, a combination of lipase-catalyzed enzymatic resolution and chemical

inversion techniques were successfully used (Tanaka, et al., 2002).

24

Flavor and aroma industry

Producing flavor esters by extraction from plant sources and fermentation is time-

consuming, expensive and restricted to the supply of natural materials. One alternative

is the use of lipases to catalyze the production of flavor and fragrance (González-

Navarro and Braco. 1998; González-Navarro and Braco. 1998; Claon and Akoh. 1993;

Karrachaabouni, et al., 1996). Among the several examples of lipases in the

development of food flavor is the use of the Mucor miehei lipase to catalyze

esterification of citronellol and geraniol with short-chain fatty acids (Laboret and Perraud.

1999). Fungal and pancreatic lipases were used to enhance lipolysis reactions and the

development of piquant flavor and sharp odor in Idiazabal cheese (a cheese made from

raw ewes' milk) (Barron, et al., 2004).

(−)-Menthol is a component of peppermint oil and is produced on industrial scale by

optical resolution of (±)-menthol. (−)-Menthol and its esters are more important from the

industrial point of view than (±)-menthol. (−)-Menthol, because of its cooling and

refreshing effects, is an important fragrance and flavor compound that is used largely in

cosmetics, toothpaste, chewing gums, cigarettes, sweets and medicines. (−)-menthol

was synthesized by enantioselective transesterification of (±)-menthol using

Burkholderia cepacia lipase (Athawale, et al., 2001).

Food industry

Fats and oil modification is one of the hot areas in food processing industry, tailored

vegetable oils with nutritionally important structured triacylglycerols and altered

physicochemical properties could have a big market value. It is the main interest of

members in the European Federation for the Science and Technology of Lipids

25

(http://www.eurofedlipid.org/). Lipases, especially microbial lipases, which are regio-

specific, are exploited for retailoring of vegetable oils (Gupta, et al., 2003). A good

example of processing fat is the use of immobilized lipases to catalyze batch-directed

interesterification of tallow, resulting in oleins containing significantly higher levels of

unsaturated fatty acids than obtained by fractionation without lipase (MacKenzie and

Stevenson, 2000).

Several lipases are used to catalyze the synthesis of structured lipids. In order to

produce one kind of reduced-calorie structured lipids, different lipases were used to

catalyze the incorporation of short-chain fatty acids (acetic, propionic, and butyric acids)

into triolein (Tsuzuki, 2005).

Improving lipases for efficient applications

For biocatalysts in general and lipases specifically, turn over rates and stability

under the application conditions are the major issues that need to be improved. Lipases

with potential application for detergent industry have to be stable against proteolytic

action, thermostable, alkalistable, stable against oxidative compounds and detergent

ingredients and preferred to have low substrate specificity. A desire for kinetic and turn

over rate changes is more likely needed in the food, chemical and pharmaceutical areas.

The performance of an enzyme that is active in one given reaction is not always

sufficient for its application in an industrial process. Consequently, there are various

chemical, physical and genetic modifications strategies proposed and applied to

enhance the properties and discover the principal suitability of a given lipase

(Bornscheuer, et al., 2002; Villeneuve, et al., 2000). These strategies are also

applicable to other biocatalysts. Some of these strategies include immobilization of the

26

lipases in a defined region, that is enclosed by material barrier for physical separation of

the enzyme from the reaction medium, and at the same time permeable to reactants

and products, such as microencapsulation using nanofibrous membranes (Ye, et al.,

2006), reverse micelles (Carvalho and Cabral, 2000), or it could be achieved by

attachment to a carrier either covalently (Yemul and Imae, 2005), by hydrophobic

interactions (Dosanjh, and Kaur, 2002), by ion exchange adsorption or by cross linking

(Kartal. and Kilinc. 2006; Kilinc, et al., 2006). Mineral support such as glass beads

(Marlot, et al., 1985), silica (Wisdom, et al., 1985; Ivanov and Schneider, 1997.) and

alumina (Brady, et al., 1986) have also been used. However, the most recent used

supports are ion exchange resins, biopolymers (Gitlesen, et al., 1997; Gray, et al., 1990)

and celite (Svensson, et al., 1990; Kang, et al., 1988; Ivanov and Schneider, 1997;

Kilara and Shahani1, 1977). Immobilization often stabilizes a biocatalyst and facilitates

downstream processing by easy separation; in addition it allows the repeated use of the

enzyme and thus significant reduction in the operating costs (Balcão, et al., 1996, Kilara

and Shahanil, 1977).

Protein engineering by rational protein design using recombinant DNA technologies

allows the amino sequence of the lipase to be changed so that it acquires different

properties (specificity, selectivity, and stability) that can better fit a particular application.

Liebeton et al. has dramatically enhanced the enantioselectivity of the lipase from

Pseudomonas aeruginosa by performing successive rounds of random mutagenesis

using error-prone (ep)-PCR (Liebeton, et al., 2000). A combination of error-prone PCR

with DNA shuffling was effective to produce a lipase variant of Pseudomonas

aeruginosa with complete enantioselectivity inversion (Zha, et al., 2001). A combination

27

of error-prone- PCR and cell surface display were successfully used to enhance the

activity of Rhizopus oryzae lipase for both hydrolysis and esterification reactions

(Shiraga, et al., 2005). Another study focused on exploring the effects of altered active

site accessibility and protein backbone flexibility on the catalytic performance of lipase B

from Candida antarctica. Qian and Lutz (2005) employed circular permutation that is the

relocating of the protein's N- and C-termini; kinetic analysis indicated that a majority of

enzyme variants either retained or surpassed wild-type activity on a series of standard

substrates.

To examine the mutational effect on a protein based on only five residues for

example, 3,200,000 (=205) variants should be theoretically covered. Kato and

coworkers (2005) have established a strategy for exploring functional proteins

associated with computational analysis by using fuzzy neural network (FNN). FNN is a

type of artificial neural network, which automatically constructs complex model

structures by learning the hidden relationship between input and output data, and it

functions as a predictor. They used this approach to screen lipases with inverted

enantioselectivity, from the (S)-form substrate to the (R)-form substrate, and they

successfully reversed the enantioselectivity of the wild-type lipase of Burkholderia

cepacia KWI-56 from the (S)-configuration to the (R) form for substrate p-nitrophenyl 3-

phenylbutyrate.

Lipases from extreme microorganisms

Extremophiles includes organisms able to thrive at extremes of temperature,

pressure, low water activity, salinity, acidity, alkalinity, radiation and cmpination thereof.

28

However, some extremophiles can also be isolated from non extremophilic

environments (Wiegel, 1998; Engle et al., 1996).

As a result, extremophiles have the potential to produce uniquely valuable

biocatalysts that function under conditions in which, their usually non-extremophilic

counterparts could not. A prominent example is the DNA polymerase I (Taq polymerase)

from the bacterium Thermus aquaticus and Pfu polymerase from the archaeon

Pyrococcus fusarium, which are predominantly used in polymerase chain reactions

(Kaledin, et al., 1980; Innis, et al., 1988). Here, we summarize the properties of lipases

from organisms that grow at two extreme environments, i.e. low (psychrophiles) and

high (thermophiles) temperatures. Other extreme conditions appear presently to be of

less interest.

Psychrophilic lipases

The detergent industry has made a shift to seek lipases from psychrophilic

organisms, since washing at low temperature will save energy and lower the cost, and

make it affordable to developing countries especially India and China. The search for a

lipase producing psychrophilic bacteria was started in the early 70’s, by isolating lipolytic

Acinetobacter sp. (Breuil and Kushner, 1975). Several psychrotolerant lipolytic

Moraxella species were subsequently isolated from the antarctic sea water, they all

produce lipases that have high activity, but not optimum, in the temperature range of 0

to 20°C (Feller, et al., 1990). Consequently, three lipases genes from Moraxella TA144

were sequenced and cloned in E. coli (Feller, et al., 1990, 1991). This was followed by

isolation of many other psychrophilic lipolytic bacteria including Psychrobacter immobilis

B10 (Arpigny, et al., 1993), Pseudomonas sp.B11-1 (Choo, et al., 1998), Acinetobacter

29

calcoacetius LP009 (Pratuangdejkul and Dharmsthiti, 2000), Arthrobacter flavus

CMS19YT (Reddy, et al., 2000), Psychrobacter okhotskensis (Yumoto, et al., 2003),

and the psychrotolerant bacterium Corynebacterium paurometabolum MTCC 6841

(Joshi, et al., 2006).

One factor that contributes to the efficient catalytic activity of psychrophilic enzymes

at lower temperature is the increase in the enzyme’s domains flexibility. In contrast to

thermophilic and mesophilic proteins, the number of salt bridges, ionic interactions,

hydrogen bonds, hydrophobic and/or inter subunits interactions are usually lower in

psychrophilic proteins (Bentahir, et al., 2000; Kim, et al., 1999; Fields, 2001; Gerday, et

al., 1997).

Thermophilic lipases

The most investigated enzymes from extremophiles are those from thermophiles

(Wiegel, 1998). These enzymes are generally the most stable at high temperature and

stable in organic solvents (Pantazaki, et al., 2002; Ejima, et al., 2004; Fucinos, et al.,

2005; Li and Zhang, 2005). Although there are some enzymes from mesophilic sources

that withstand elevated temperatures, such cases are rare (e.g. B. cepacia lipase).

Thermophilic enzymes serve an excellent models for understanding protein stability and

carry significant potential for biotechnology, for instance, factors that can contribute to

the high thermostability of a given enzyme include changes in amino acid residues,

increased salt-bridge content, reductions in cavity size, increased hydrophobic

interactions and changes in solvent-exposed surface areas (Demirjian, et al., 2001;

Adams and Kelly, 1998; Eichler, 2001).

30

Table 1.4 lists examples of thermophilic lipases. One of the most interesting lipases

was isolated from Thermosyntropha lipolytica, which is the subject of this study. This

strain constitutively produces two lipases that show maximal activity at 96 °C depending

on the assay buffer and retain 50% activity after 20 hours incubation at 75 °C. The only

other lipase with such high thermal activity and stability was surprisingly found in the

mesophilic Burkholderia cepacia (Rathi, et al., 2000). However, comparing the physical

properties of these lipases is quite difficult due to the differences in the assay substrates

and procedure. Most of the lipases that are commercially adopted by many companies

are from Pseudomonas species and this is due to the fact that these lipases are the first

to be isolated, cloned and characterized, their 3-D structures among the earliest to be

revealed along with well established biochemical and genetic data. Moreover, most of

these lipases were gone through a variety of molecular modifications to produce

effective enzymes with more desirable characteristics (Jaeger and Reetz. 1998, Gupta,

et al., 2004).

The objectives and significance of this work.

Numerous bacterial lipases have been isolated and purified, and/or their genes

cloned and expressed to high levels (Table 1.5). As is obvious from the above review

little is known about lipases from anaerobes and particularly anaerobic thermophiles.

Because of their biotechnological potential, it is of great interest to determine their

stereo-specificities, fatty acid specificities, or alcohol moiety preferences. Thus, this

study elucidated the properties of two extremely thermostable lipases from

Thermosyntropha lipolytica. This is an anaerobic thermophilic bacterium that degrades

lipids in syntrophic relationship with a hydrogen utilizing microorganism (Svetlitshnyi, et

31

al., 1996). Chapter 2 of this dissertation reports on the isolation and purification of two

extracellular lipases, LipA and LipB, and their general characterization including their

temperature and pH optima, substrate specificity and regio-stereospecificity. Chapter 3

discusses the effect of anionic, cationic and non ionic detergents on the activity and

stability of the two lipases as well as the biochemical and kinetic studies in the absence

and presence of SDS. Chapter 4 covers the oligomerization behavior of both lipases as

noticed by gel filtration and native gradient gel electrophoresis, and the effect of

oligomerization on thermostability. Chapter 5 shows the ability of both enzymes to carry

out the reverse reaction and to catalyze the synthesis of different fatty alcohol esters

and glyceride moieties, and characterization in different organic solvents.

32

References

Abousalham, A., Nari, J., Teissere, M., Ferte, N., Noat, G., and Verger, R. (1997).

Study of fatty acid specificity of sunflower phospholipase D using

detergent/phospholipid micelles Eur. J. Biochem. 248, 374-379.

Adams, M. W., and Kelly, R. M. (1998). Finding and using hyperthermophilic enzymes.

Trends Biotechnol. 16, 329–332.

Akimoto, T., and Nagase, Y. (2003). Novel transdermal drug penetration enhancer:

synthesis and enhancing effect of alkyldisiloxane compounds containing

glucopyranosyl group. J. Control Release. 88, 243-252.

Al-Zuhair, S. (2005). Production of biodiesel by lipase-catalyzed transesterification of

vegetableoils: a kinetics study. Biotechnol. Prog. 21, 1442-1448.

Arcos, J. A., Hill, C. G. Jr., and Otero, C. 2001. Kinetics of the lipase-catalyzed

synthesis of glucose esters in acetone. Biotechnol. Bioeng. 73, 104-110.

Arpigny, J. L Feller, G. and C. Gerday. 1993. Cloning, sequence and structural

features of a lipase from the antarctic facultative psychrophile Psychrobacter

immobilis B10. Biochim. Biophys. Acta. 1171, 331-333.

Arpigny, J. L., and Jaeger, K-E. (1999). Bacterial lipolytic enzymes: classification and

properties. Biochem. 343, 177–183.

Athawale, V., Manjrekar, N., and Athawale, M. (2001). Enzymatic synthesis of chiral

menthyl methacrylate monomer by Pseudomonas cepacia lipase catalysed

resolution of (±)-menthol. J. Mol. Catal. B: Enzymatic. 16, 169–173.

Balcão, V. M., Paiva, A. L., and Malcata, F. X. (1996). Bioreactors with immobilized

lipases: State of the art. Enzyme Microb. Technol. 18, 392–416.

33

Barron, L. J., Hernandez, I., Bilbao, A., Flanagan, C. E., Najera, A. I., Virto, M., Perez-

Elortondo, F. J., Albisu, M., and de Renobales, M. (2004). Changes in lipid

fractions and sensory properties of Idiazabal cheese induced by lipase addition. J.

Dairy. Res. 71, 372-379.

Bell, P. J. L., Nevalainen, H., Morgan, H. W., and Bergquist. P. L. (1999). Rapid

cloning of thermoalkalophilic lipases from Bacillus sp. Using PCR. Biotech. Lett.

21, 1003-1006.

Bentahir, M., Feller, G., Aittaleb, M., Lamotte-Brasseur, J., Himri, T., Chessa, J. P.,

and Gerday, C. (2000). Structural, kinetic, and calorimetric characterization of the

cold-active phosphoglycerate kinase from the Antarctic Pseudomonas sp.

TACII18. J. Biol. Chem. 275, 11147–11153.

Blow, D. M., Birktoft, J. J., and Hartley, B. S. (1969). Role of a buried acid group in the

mechanism of action of chymotrypsin. Nature. 221, 337-340.

Bourne, P. C., Isupov, M. N., and Littlechild, J. A. (2000). The atomic-resolution

structure of a novel bacterial esterase. Struct. Fold. Des. 8, 143–151.

Bourne, Y., Hasper, A. A., Chahinian, H., Juin, M., De Graaff, L. H., and Marchot, P.

(2004). Aspergillus niger protein EstA defines a new class of fungal esterases

within the alpha/beta hydrolase fold superfamily of proteins. Structure (Camb). 12,

677-87.

Bornscheuer, U. T., Bessler, C., Srinivas, R., and Krishna, S. H. (2002). Optimizing

lipases and related enzymes for efficient application. Trends Biotechnol. 20, 433-

7.

34

Brady, L., Brzozowski, A. M., Derewenda, Z. S., Dodson, E., Dodson, G., Tolley, S.,

Turkenburg, J. P., Christiansen, L., Huge-Jensen, B., Norskov, L., Thim, L., and

Menge, U. (1990). A serine protease triad forms the catalytic centre of a

triacylglycerol lipase. Nature. 343,767-770.

Brady, C., Metcalfe. L., Slaboszewski, D., and Frank, D. (1986). Hydrolysis of liquid

fats using lipase adsorbed on porous substrate of untreated microporous

polyolefin, from aq. medium, using fixed bed or stirred reactors. US Patent No.

4,629,742.

Breuil, C., and Kushner D. J. (1975). Lipase and esterase formation by psychrophilic

and mesophilic Acinetobacter species. Can. J. Microbiol. 21, 423-433.

Broadrup, R. L., Sundar, H. M., and Swindell, C. S. (2005). Total synthesis of 12,13-

desoxyepothilone B (Epothilone D). Bioorg. Chem. 33, 116-133.

Bryant, M. P., Campbell, L. L., Reddy, C. A., Crabill, M. R. (1997). Growth of

Desulfovibrio in lactate or ethanol media low in sulfate in association with H2-

utilizing methanogenic bacteria. Appl. Environ. Microbiol. 33,1162-1169.

Bryant, M. P., Wolin, E. A., Wolin, M. J., and Wolfe, R. S. 1967. Methanobacillus

omelianskii, a symbiotic association of two species of bacteria. Arch. Mikrobiol.

59, 20-31.

Brzozowski A. M., Derewenda, U., Derewenda, Z. S., Dodson, G. G., Lawson, D. M.,

Turkenburg, J. P., Bjorkling, F., Huge-Jensen, B., Patkar, S. A., and Thim, L.

(1991). A model for interfacial activation in lipases from the structure of a fungal

lipase-inhibitor complex. Nature. 351, 491 494.

35

Carvalho, C. M., and Cabral J. M. (2000). Reverse micelles as reaction media for

lipases. Biochimie. 82,1063-85.

Chahinian, H., Ali, Y. B., Abousalham, A., Petry, S., Mandrich, L., Manco, G., Canaan,

S., and Sarda, L. (2005). Substrate specificity and kinetic properties of enzymes

belonging to the hormone-sensitive lipase family: comparison with non-lipolytic

and lipolytic carboxylesterases. Biochim Biophys Acta. 1738, 29-36.

Chahinian, H., Nini, L., Boitard, E., Dubes, J. P., Comeau, L. C., and Sarda, L. (2002).

Distinction between esterases and lipases: a kinetic study with vinyl esters and

TAG. Lipids. 37, 653-662.

Chihara-Siomi, M., Yoshikawa, K., Oshima-Hirayama, N., Yamamoto, K., Sogabe, Y.,

Nakatani, T., Nishioka, T., and Oda, J. (1992). Purification, molecular cloning,

and expression of lipase from Pseudomonas aeruginosa. Arch. Biochem.

Biophys. 296, 505-513.

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 Microbiol. Lett. 186, 235-238.

Choo, D. W., Kurihara, T., Suzuki, T., Soda K., and Esaki, N. (1998). A cold-adapted

lipase of an Alaskan psychrotroph, Pseudomonas sp. strain B11-1: gene cloning

and enzyme purification and characterization. Appl Environ Microbiol 64, 486–

491

Claon, P. A., and Akoh, C. C. (1993). Enzymatic synthesis of geraniol and citronellol

esters by direct esterification in n-hexane. Biotechnol. Lett. 15, 1211-1215.

36

Contreras, J. A., Karlsson, M., Osterlund, T., Laurell, H., Svensson, A., and Holm, C.

(1996). Hormone-sensitive lipase is structurally related to acetylcholinesterase,

bile salt-stimulated lipase, and several fungal lipases. Building of a three-

dimensional model for the catalytic domain of hormone-sensitive lipase. J. Biol.

Chem. 271, 31426-30.

Cygler, M., and Schrag, J. D. (1997). Structure as basis for understanding interfacial

properties of lipases. Methods Enzymol. 284, 3–27.

Dartois, V., Baulard, A., Schanck, K., and Colson, C. (1992). Cloning, nucleotide

sequence and expression in Escherichia coli of a lipase gene from Bacillus

subtilis 168. Biochim. Biophys. Acta. 1131, 253-60.

Dartois, V., Coppee, J. Y., Colson, C. and Baulard, A. (1994). Genetic analysis and

overexpression of lipolytic activity in Bacillus subtilis. App. Env. Microbiol. 60,

1670-1673.

Deems, R. A., Eaton, B. R., and Dennis, E. A. (1975). Kinetic analysis of

phospholipase A2 activity toward mixed micelles and its implications for the study

of lipolytic enzymes. J. Biol. Chem. 250, 9013-9020.

Demirjian, D. C., Moris-Varas, F., and Cassidy, C. S. (2001). Enzymes from

extremophiles. Curr. Opin. Chem. Biol. 5, 144-51.

Derewenda, U., Brzozowski, A. M., Lawson, D. M., and Derewenda, Z. S. (1992).

Catalysis at the interface: the anatomy of a conformational change in a

triglyceride lipase. Biochemistry. 31, 1532 1541.

37

Dharmsthiti, S., and Luchai, S. (1999). Production, purification and characterization of

thermophilic lipase from Bacillus sp. THL027. FEMS Microbiol. Lett. 179, 241-

246.

Dong, H., Gao, S., Han, S., and Cao, S. (1999). Purification and characterization of a

Pseudomonas sp. lipase and its properties in non-aqueous media. Biotechnol.

Appl. Biochem. 30, 251-256.

Dosanjh, N. S., and Kaur, J., (2002). Immobilization, stability and esterification studies

of a lipase from a Bacillus sp. Biotechnol Appl Biochem. 36, 7-12.

Du, W., Xu, Y. Y., Zeng, J., and Liu, D. H. (2004). Novozym 435-catalysed

transesterification of crude soya bean oils for biodiesel production in a solvent-

free medium. Biotechnol. Appl. Biochem. 40, 187-90.

Duong, F., Soscia, C., Lazdunski, A., and Murgier, M. (1994). The Pseudomonas

fluorescens lipase has a C-terminal secretion signal and is secreted by a three-

component bacterial ABC-exporter system. Mol. Microbiol. 11, 1117-26.

Eggert, T., Pencreac'h, G., Douchet, I., Verger, R., and Jaeger, K-E. 2000. A novel

extracellular esterase from Bacillus subtilis and its conversion to a

monoacylglycerol hydrolase. Eur. J .Biochem. 267, 6459-69.

Eichler, J. (2001). Biotechnological uses of archaeal extremozymes. Biotechnol. Adv.

19, 261-278.

Ejima, K., Liu, J., Oshima, Y., Hirooka, K., Shimanuki, S., Yokota, Y., Hemmi, H.,

Nakayama, T., and Nishino, T. (2004). Molecular cloning and characterization of

a thermostable carboxylesterase from an archaeon, Sulfolobus shibatae

38

DSM5389: Non-linear kinetic behavior of a hormone-sensitive lipase family

enzyme. J. Biosci. Bioeng. 98, 445-451.

Farooqui, A. A. Taylor, W. A. and L.A. Horrocks. 1987. Phospholipases,

lysophospholipases and lipases and their involvement in various diseases.

Neurochem. Pathol. 7: 99-128.

Feller, G., Thiry, M., Arpigny, J. L., and Gerday, C. (1991). Cloning and expression in

Escherichia coli of three lipase-encoding genes from the psychrotrophic antarctic

strain Moraxella TA144. Gene. 102,111-5.

Feller, G., Thiry, M., Arpigny, J. L., Mergeay, M., and Gerday, C. (1990). Lipases from

psychrotrophic antarctic bacteria. FEMS Micriobiol. Lett. 66, 239-244.

Feller, G., Thiry, M., and Gerday, C. (1991). Nucleotide sequence of the lipase gene

lip2 from the antarctic psychrotroph Moraxella TA144 and site-specific

mutagenesis of the conserved serine and histidine residues. DNA Cell Biol. 10,

381-388.

Feller, G., Thiry, M., and Gerday, C. (1991). Nucleotide sequence of the lipase gene

lip3 from the antarctic psychotroph Moraxella TA144. Biochim Biophys Acta.

1088, 323-324.

Feller, G Thiry, M., and Gerday, C. (1990). Sequence of a lipase gene from the

antarctic psychrotroph Moraxella TA144. Nucleic Acids Res. 18, 6431.

Ferrato, F., Carriere, F., Sarda, L., and R. Verger. (1997) A critical reevaluation of the

phenomenon of interfacial activation. Methods Enzymol. 286, 327-347.

Fields, P. A. (2001). Protein function at thermal extremes: balancing stability and

flexibility. Comp. Biochem. Physiol. 129, 417–431.

39

Fischer, M., and Pleiss, J. (2003). The Lipase Engineering Database: a navigation and

analysis tool for protein families. Nucleic Acids Res. 31:319-21.

Flores, M. V. and P. J. Halling. (2002). Full model for reversible kinetics of lipase-

catalyzed sugar-ester synthesis in 2-methyl 2-butanol. Biotechnol. Bioeng. 78,

795-801.

Fucinos, P. Dominguez, A. Sanroman, M. A., Longo, M. A., Rua, M. L., and Pastrana,

L. (2005). Production of thermostable lipolytic activity by Thermus species.

Biotechnol. Prog. 21, 1198-205.

Fukuda, H., Kondo, A., and Noda, H. (2001). Biodiesel fuel production by

transesterification of oils. Biosci. Bioeng. 92, 405-416.

Gerday, C., Aittaleb, M., Arpigny, J. L., Baise, E., Chessa, J. P., Garsoux, G.,

Petrescu, I., and Feller, G. (1997). Psychrophilic enzymes: a thermodynamic

challenge. Biochim. Biophys. Acta. 1342, 119-131.

Gitlesen, T., Bauer, M., and Adlercreutz, P. (1997). Adsorption of lipase on

polypropylene powder. Biochem. Biophys. Acta. 1345, 188-196.

Godfry, T., and West, S. (1996). “Industrial Enzymology. The application of Enzymes

in industry.” Nature press, New Yok.

González-Navarro, H., and Braco, L. (1998). Lipase-enhanced activity in flavour ester

reactions by trapping enzyme conformers in the presence of interfaces.

Biotechnol Bioeng. 59, 122-127.

Gotz, F., Verheij, H. M., and Rosenstein, R. (1998). Staphylococcal lipases: molecular

characterization, secretion, and processing. Chem. Phys. Lipids. 93, 15-25.

40

Gray, C. J., Narang, J. S., and Barker, S. A. (1990). Immobilization of lipase from

Candida cylindracea and its use in the synthesis of menthol esters by

transesterification. Enzyme Microb. Technol. 12, 800-807

Gupta, R., Gupta, N., and Rathi, P. (2004). Bacterial lipases: an overview of

production, purification and biochemical properties. Appl. Microbiol. Biotechnol.

64, 763-781.

Gupta, R., Rathi, P., and Bradoo, S. (2003). Lipase mediated up gradation of dietary

fats and oils. Crit. Rev. Food Sci. Nutr. 43, 635-644.

Gu, J., Ruppen, M. E., and Cai, P. (2005). Lipase-catalyzed regioselective

esterification of rapamycin: synthesis of temsirolimus (CCI-779). Org. Lett. 7,3

945-948.

Hata, Y., Matsuura, Y., Tanaka, N., Kakudo, M., Sugihara, A., Iwai, M., and Tsujisaka,

Y. (1979). Low resolution crystal structure of lipase from Geotrichum candidum

(ATCC34614). J. Biochem. 86, 1821-1827.

Holmquist, M. (2000). Alpha/Beta-hydrolase fold enzymes: structures, functions and

mechanisms. Curr. Protein Pept. Sci. 1, 209-235.

Hjorth, A., Carriere, F., Cudrey, C., Woldike, H., Boel, E., Lawson, D. M., Ferrato, F.,

Cambillau, C., Dodson, G. G., Thim, L., and Verger, R. (1993). A structural

domain (the lid) found in pancreatic lipases is absent in the guinea pig (phospho)

lipase. Biochemistry. 32, 4702-4707.

Innis, M. A., Myambo K. B., Gelfand D. H., and Brow M. A. (1988). DNA sequencing

with Thermus aquaticus DNA polymerase and direct sequencing of polymerase

chain reaction-amplified DNA. Proc. Natl. Acad. Sci. U S A. 85, 9436-9440.

41

Iso, M., Chen, B., Eguchi, M., Kudo, T., and Shrestha, S. (2001). Production of

biodiesel fuel from triglycerides and alcohol using immobilized lipase. J. Mol.

Catal. B: Enzymatic. 16, 53–58.

Ivanov, A. E. and M.P. Schneider. 1997. Methods for the immobilization of lipases and

their use for ester synthesis. J. Mol. Catal. B Enzym. 3, 303–309.

Jackson, B. E., Bhupathiraju, V. K., Tanner, R. S., Woese, C. R., and McInerney, M. J.

(1999). Syntrophus aciditrophicus sp. nov., a new anaerobic bacterium that

degrades fatty acids and benzoate in syntrophic association with hydrogen-using

microorganisms. Arch. Microbiol. 171, 107-14.

Jaeger K. E., and Eggert, T. (2002). Lipases for biotechnology. Curr. Opin. Biotechnol.

13, 390-397.

Jaeger K. E., and Reetz, M. T. (1998). Microbial lipases form versatile tools for

biotechnology. Trends Biotechnol. 16, 396-403.

Jaeger, K-E, Dijkstra, B. W., and Reetz, M. T. (1999). Bacterial biocatalysts: molecular

biology, three-dimensional structures, and biotechnological applications of

lipases. Annu. Rev. Microbiol. 53, 315–351.

Jaeger, K-E., Ransac, S., Dijkstra, B. W., Colson, C., Van Heuvel, M., Misset, O. 1994.

Bacterial lipases. FEMS Microbiol. Rev. 15: 29-63.

Jeong, S. T., Kim, H. K., Kim, S. J., Chi, S. W., Pan, J. G., Oh, T. K., and Ryu, S. E.

(2002). Novel zinc-binding center and a temperature switch in the Bacillus

stearothermophilus L1 lipase. J. Biol. Chem. 277, 17041-17047.

42

Joshi G. K., Kumar S., Tripathi B. N., Sharma V. (2006). Production of Alkaline Lipase

by Corynebacterium paurometabolum, MTCC 6841 Isolated from Lake

Naukuchiatal, Uttaranchal State, India. Curr Microbiol. 52, 354-358.

MacKenzie, A. D., and Stevenson, D. E. (2000). Production of high-oleic acid tallow

fractions using lipase-catalyzed directed interesterification, using both batch and

continuous processing. Enzyme Microb. Technol. 27, 302-311.

Kaledin A. S., Sliusarenko A. G., and Gorodetskii S. I. (1980). Isolation and properties

of DNA polymerase from extreme thermophylic bacteria Thermus aquaticus YT-1.

Biokhimiia. 45, 644-651.

Kang, S. T., and Rhee, J. S. (1988). Effect of water on hydrolysis of olive oil by

immobilized lipase in reverse phase system. Biotechnol. Lett. 10, 341-346.

Karrachaabouni, M., Pulvin, S., Touraud, D., and Thomas, D. (1996). Enzymatic

synthesis of geraniol esters in a solvent free system by lipases. Biotechnol. Lett.

18, 1083-1088.

Kartal, F., and Kilinc A. (2006). Immobilization of pancreatic lipase on polyvinyl alcohol

by cyanuric chloride. Prep Biochem Biotechnol. 36, 139-151.

Kato, R., Nakano, H., Konishi, H., Kato, K., Koga, Y., Yamane, T., Kobayashi, T., and

Honda, H. (2005). Novel strategy for protein exploration: high-throughput

screening assisted with fuzzy neural network. J. Mol. Biol. 351, 683-692.

Kazlauskas, R. J. (1994). Elucidating structure-mechanism relationships in lipases:

prospects for predicting and engineering catalytic properties. Trends Biotech. 12,

464-472.

43

Kilara, A., and Shahanil, C. R. C. (1977). The use of immobilized enzymes in the food

industry. Crit. Rev. Food Sci. Nutr. 12, 161-198.

Kim, H. K., Choi, H. J., Kim, M. H., Sohn, C. B., and Oh, T. K. (2002). Expression and

characterization of Ca (2+)-independent lipase from Bacillus pumilus B26.

Biochim. Biophys. Acta. 1583, 205-212.

Kim, S. Y., Hwang, K. Y., Kim, S. H., Sung, H. C., Han Y. S., and Cho, Y. (1999).

Structural basis for cold adaptation. Sequence, biochemical properties, and

crystal structure of malate dehydrogenase from a psychrophile Aquaspirillium

actricum. J. Biol. Chem. 274, 11761–11767.

Kim, H. K., Park, S. Y., Lee, J. K. and T. K. OH. 1998. Gene cloning and

characterization of thermostable lipase from Bacillus stearothermophilus L1.

Biosci. Biotechnol. Biochem. 62, 66-71.

Kim, K. K., Song, H. K., Shin, D. H., Hwang, K. Y., Choe, S., Yoo, O. J., and Suh, S.

W. (1997). Crystal structure of carboxylesterase from Pseudomonas fluorescens,

an alpha/beta hydrolase with broad substrate specificity. Structure. 5, 1571–1584.

Kim, K. K., Song, H. K., Shin, D. H., Hwang, K. Y., and Suh, S. W. (1997). The crystal

structure of a triacylglycerol lipase from Pseudomonas cepacia reveals a highly

open conformation in the absence of a bound inhibitor. Structure. 5, 173-185.

Kilinc, A., Teke, M., Onal, S., Telefoncu, A. (2006). Immobilization of pancreatic lipase

on chitin and chitosan. Prep Biochem Biotechnol. 36, 153-163.

Kitaura, S., Suzuki, K., and Imamura, S. (2001). Monoacylglycerol lipase from

moderately thermophilic Bacillus sp. strain H-257: molecular cloning, sequencing,

and expression in Escherichia coli of the gene. J. Biochem. 129, 397-402.

44

Kojima Y., Kobayashi, M., and Shimizu, S. (2003). A novel lipase from Pseudomonas

fluorescens HU380: gene cloning, overproduction, renaturation-activation, two-

step purification, and characterization. J Biosci. Bioeng. 96, 242-249.

Kojima and Shimizu. 2003. Purification and characterization of the lipase from

Pseudomonas fluorescens HU380. J. Biosci. Bioeng. 96, 219-226.

Kojima, Y., Yokoe, M., and Mase, T. (1994). Purification and characterization of an

alkaline lipase from Pseudomonas fluorescens AK102. Biosci. Biotechnol.

Biochem. 58, 1564-1568.

Kordel, M., Hofmann, B., Schomburg, D., and Schmid, R. D. (1991). Extracellular

lipase of Pseudomonas sp. strain ATCC 21808: purification, characterization,

crystallization, and preliminary X-ray diffraction data. J. Bacteriol. 173, 4836-4841.

Laboret, F., and Perraud, R. (1999). Lipase-catalyzed production of short-chain acids

terpenyl esters of interest to the food industry. Appl. Biochem. Biotechnol. 82,

185-98.

Labuschagne, R. B., Tonder, A. V., and Litthauer, D. (1997). Flavobacterium odoratum

lipase: isolation and characterization. Enzyme Microb. Technol. 21, 52-58.

Lang, D., Hofmann, B., Haalck, L., Hecht, H. J., Spener, F., Schmid, R. D., and

Schomburg, D. (1996). Crystal structure of a bacterial lipase from

Chromobacterium viscosum ATCC 6918 refined at 1.6 angstroms resolution. J.

Mol. Biol. 259, 704-717.

Lesuisse, E., Schanck, K., and Colson, C. (1993). Purification and preliminary

characterization of the extracellular lipase of Bacillus subtilis 168, an extremely

basic pH-tolerant enzyme. Eur. J. Biochem. 216, 155-160.

45

Liebeton, K., Zonta, A., Schimossek, K., Nardini, M., Lang, D., Dijkstra, B. W., Reetz,

M. T., and Jaeger, K-E. (2000). Directed evolution of an enantioselective lipase.

Chem. Biol. 7, 709-718.

Li, X., Tetling, S., Winkler, U. K., Jaeger, K. E., and Benedik, M. J. (1995). Gene

cloning, sequence analysis, purification, and secretion by Escherichia coli of an

extracellular lipase from Serratia marcescens. Appl. Environ. Microbiol. 61, 2674-

2680.

Li H., and Zhang, X. (2005). Characterization of thermostable lipase from thermophilic

Geobacillus sp. TW1. Protein Expr. Purif. 42, 153-159.

Lin, S. F., Chiou, C. M., Yeh, C. M., and Tsai, Y. C. (1996). Purification and partial

characterization of an alkaline lipase from Pseudomonas pseudoalcaligenes F-

111. Appl. Environ. Microbiol. 62, 1093-1095.

Liu, H. L., Hoff, B. H., Berg, T. C., and Anthonsen, T. (2001). Synthesis of the

antistroke drug lubeluzole and its enantiomer. Lipase-catalyzed resolution of

chiral building block. Chirality. 13, 135-139.

Lorowitz, W. H., Zhao, H., and Bryant, M. P. (1989). Syntrophomonas wolfei subsp.

saponavida subsp. nov., a long-chain fatty-acid-degrading, anaerobic, syntrophic

bacterium; Syntrophomonas wolfei subsp. wolfei subsp. nov.; and emended

descriptions of the genus and species. Int J Syst Bacteriol. 39, 122–126.

Lucas, K. H., and Kaplan-Machlis, B. (2001). Orlistat--a novel weight loss therapy. Ann

Pharmacother. 35, 314-328.

Mamputu, J. C, and Renier, G. (1999). Differentiation of human monocytes to

monocyte-derived macrophages is associated with increased lipoprotein lipase-

46

induced tumor necrosis factor-alpha expression and production: a process

involving cell surface proteoglycans and protein kinase C. Arterioscler. Thromb.

Vasc. Biol. 19, 1405-1411.

Malcata, F. X., Reyes, H. R., Garcia, H. S., Hill Jr., C. G., and Amundson, C. H. (1992).

Kinetics and mechanisms of reactions catalyzed by immobilized lipases. Enzyme

Microb. Technol. 14: 426-446.

Marlot, C., Langrand, G., Triantaphylides, C., and Baratti, J. (1985). Ester synthesis in

organic solvent catalyzed by lipases immobilized on hydrophilic supports.

Biotechnol. Lett. 7, 647-650.

Martinez, C., De Geus, P., Lauwereys, M., Matthyssens, G., and Cambillau, C. (1992).

Fusarium solani cutinase is a lipolytic enzyme with a catalytic serine accessible

to solvent. Nature. 356, 615-618.

McInerney, M. J., Bryant, M. P., and Pfennig, N. (1979). Anaerobic bacterium that

degrades fatty acids in syntrophic association with methanogens. Arch. Microbial.

122, 129-135.

McInerney, M. J., Mackie, R. I., and Bryant, M. P. (1981). Syntrophic association of a

butyrate-degrading bacterium and methanosarcina enriched from bovine rumen

fluid. Appl. Environ. Microbiol. 41, 826-828.

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.

Nagy, I., Banerjee, T., Tamura, T., Schoofs, G., Gils, A., Proost, P., Tamura, N.,

Baumeister, W., and De Mot. R. (2003). Characterization of a novel intracellular

47

endopeptidase of the alpha/beta hydrolase family from Streptomyces coelicolor

A3(2). J. Bacteriol. 185, 496-503.

Nardini, M., and Dijkstra, B. W. (1999). Alpha/beta hydrolase fold enzymes: the family

keeps growing. Curr Opin Struct Biol. 9, 732-737.

Nardini, M., Lang, D. A., Jaeger, K-E., and Dijkstra, B. W. (2000). Crystal structure of

Pseudomonas aeruginosa lipase in the open conformation. The prototype for

family I.1 of bacterial lipases. J. Biol. Chem. 275, 31219–31225.

Nawani, N., and Kaur, J. (1999). Purification, characterization and thermostability of

lipase from a thermophilic Bacillus sp. J33. Mol. Cell. Biochem. 206, 91-96.

Nini, L., Sarda, L., Comeau, L. C., Boitard, E., Dubes, J. P., and Chahinian, H. (2001).

Lipase-catalysed hydrolysis of short-chain substrates in solution and in emulsion:

a kinetic study. Biochim Biophys Acta. 1534, 34-44.

Noble, M. E., Cleasby, A., Johnson, L. N., Egmond, M. R., and Frenken, L. G. (1993).

The crystal structure of triacylglycerol lipase from Pseudomonas glumae reveals

a partially redundant catalytic aspartate. FEBS Lett. 331, 123-8.

Noureddini, H., Gao, X., and Philkana, R. S. (2005). Immobilized Pseudomonas

cepacia lipase for biodiesel fuel production from soybean oil. Bioresour. Technol.

96, 769-777.

Oh, B., Kim, H., Lee, J., Kang, S., and Oh, T. (1999). Staphylococcus haemolyticus

lipase: biochemical properties, substrate specificity and gene cloning. FEMS

Microbiol. Lett. 179, 385-392.

48

Ollis, D. L., Cheah, E., Cygler, M., Dijkstra, B., Frolow, F., Franken, S. M., Harel, M.,

Remington, S. J., Silman, I., Schrag, J., Sussman, J. L., Verschueren, H. K. G.,

and Goldman, A. (1992). The alpha/beta hydrolase fold. Protein Eng. 5, 197-211.

Ono, M., Suzuki, K., Tanikawa, S., and Akita, H. (2001). First synthesis of (+)- and (−)-

elvirol based on an enzymatic function. Tetrahedron Asymmetry. 12, 2597–2604.

Paiva, A. L., Balcao,V. M., and Malacta, F. X. (2000). Kinetics and mechanisms of

reactions catalyzed by immobilized lipases. Enzyme Microb. Technol. 27, 187-

204.

Pantazaki, A. A., Pritsa, A. A., and Kyriadidis, D. A. (2002). Biotechnologically relevant

enzymes from Thermus thermophilus. Appl. Microbiol. Biotechnol. 58, 1-12.

Patel R. N. (2000). Microbial/enzymatic synthesis of chiral drug intermediates. Adv.

Appl. Microbiol. 47, 33-78.

Pratuangdejkul, J., and Dharmsthiti, S. (2000). Purification and characterization of

lipase from psychrophilic Acinetobacter calcoaceticus LP009. Microbiol Res. 155,

95-100.

Qian, Z. and Lutz, S. (2005). Improving the catalytic activity of Candida antarctica

lipase B by circular Permutation. J. Am. Chem. Soc.127, 13466-13467.

Quigley, K. A., Jackson, M. L., and Haines. D. M. (2001). Hyperlipasemia in 6 dogs

with pancreatic or hepatic neoplasia: evidence for tumor lipase production. Vet.

Clin. Pathol. 30, 114-120.

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.

Biotech. Lett. 22, 495-498.

49

Raza, S., Fransson, L., and Hult, K. (2001). Enantioselectivity in Candida antarctica

lipase B: A molecular dynamics study. Protein Sci. 10, 329-338.

Reddy, G. S., Aggarwal, R. K., Matsumoto, G. I., and Shivaji, S. (2000). Arthrobacter

flavus sp. nov., a psychrophilic bacterium isolated from a pond in McMurdo Dry

Valley, Antarctica. Int. J. Syst. Evol. Microbiol. 50, 1553-1561.

Rosenstein, R., and Gotz, F. (2000). Staphylococcal lipases: biochemical and

molecular characterization. Biochimie. 82, 1005-1014.

Roy, F., Samain, E., Dubourguier, H. C., and Albagnac, G. (1986). Syntrophomonas

sapovorans sp. nov., a new obligately proton reducing anaerobe oxidizing

saturated and unsaturated long chain fatty acids. Arch Microbiol 145, 142–147.

Rua Luisa, M., Schmidt-Dannert, C., Wahl, S., Sprauer, A., and Schmid R. D. (1997).

Thermoalkalophilic lipase of Bacillus thermocatenulatus large-scale production,

purification and properties: aggregation behaviour and its effect on activity. J.

Biotech. 56, 89-102.

Ruiz, C., Blanco, A., Pastor, F. I., and Diaz, P. (2002). Analysis of Bacillus megaterium

lipolytic system and cloning of LipA, a novel subfamily I.4 bacterial lipase. FEMS

Microbiol. Lett. 217, 263-267.

Ruiz, C. Javier Pastor, F. I. and Diaz P. (2003). Isolation and characterization of

Bacillus sp. BP-6 LipA, a ubiquitous lipase among mesophilic Bacillus species.

Lett. Appl. Microbiol. 37, 354-359.

Sarda, L., and Desnuelle, P. (1958). Action of pancreatic lipase on emulsified esters.

Biochim. Biophys. Acta 30, 513-521.

50

Sayari, A., Agrebi, N., Jaoua, S., and Gargouri Y. (2001). Biochemical and molecular

characterization of Staphylococcus simulans lipase. Biochimie. 83, 863-871.

Schink, B. (1997). Energetics of syntrophic cooperation in methanogenic degradation.

Microbiol. Mol. Biol. Rev. 61, 262-280.

Schrag, J. D., and Cygler, M. (1997). Lipases and alpha/beta hydrolase fold. Methods

Enzymol. 284, 85-107.

Schrag, J. D., Li, Y., Cygler, M., Lang, D., Burgdorf, T., Hecht, H. J., Schmid, R.,

Schomburg, D., Rydel, T. J., Oliver, J. D., Strickland, L. C., Dunaway, M., Larson,

S. B., Day, J., and McPherson, A. (1997). The open conformation of a

Pseudomonas lipase. Structure. 5, 187–202.

Schrag, J. D., Li, Y., G., Wu, S., and Cygler, M. (1991). Ser-His-Glu triad forms the

catalytic site of the lipase from Geotrichum candidum. Nature. 351, 761-764.

Schmidt-Dannert, C. (1999). Recombinant microbial lipases for biotechnological

applications. Bioorg. Med. Chem. 7, 2123-2130.

Schmidt-Dannert ,C., Sztajer, H., Stocklein, W., Menge, U., and schmid, R. D. (1994).

Screening, purification and properties of a thermophilic lipase from Bacillus

thermocatenulatus. Biochim. Biophys. Acta. 1214, 43-53.

Sehanputri, P. S., and Hill Jr., C. G. (2000). Biotechnology for the production of

nutraceuticals enriched in conjugated linoleic acid: II. Multiresponse kinetics of

the hydrolysis of corn oil by a Pseudomonas sp. lipase immobilized in a hollow-

fiber reactor. Biotechnol Bioeng. 69, 450-456.

Sekiguchi, Y., Kamagata, Y., Nakamura, K., Ohashi, A., and Harada, H. (2000).

Syntrophothermus lipocalidus gen. nov., sp. nov., a novel thermophilic,

51

syntrophic, fatty-acid-oxidizing anaerobe which utilizes isobutyrate. Int J Syst

Evol Microbiol. 50, 771–779.

Shabtai, Y., and Daya-Mishne, N. (1992). Production, purification, and properties of a

lipase from a bacterium (Pseudomonas aeruginosa YS-7) capable of growing in

water-restricted environments. Appl. Environ. Microbiol. 58, 174-180.

Shiraga, S., Kawakami, M., Ishiguro, M., and Ueda, M. (2005). Enhanced reactivity of

Rhizopus oryzae lipase displayed on yeast cell surfaces in organic solvents:

potential as a whole-cell biocatalyst in organic solvents. Appl Environ Microbiol.

71, 4335-4338.

Shaw, C. C., Sellstedt, J., Noureldin, R., Cheal, G. K., and Fortier, G. (2001) U.S.

Patent 6277983.

Simons, J. W., Adams, H., Cox, R. C., Dekker, N., Gotz, F., Slotboom, A. J., and

Verheij, H. M. (1996). The lipase from Staphylococcus aureus. Expression in

Escherichia coli, large-scale purification and comparison of substrate specificity

to Staphylococcus hyicus lipase. Eur. J. Biochem. 242, 760-769.

Simons, J., Boots, J. P., Slotboom, A. J., and Verheij, H. M. (1997). The

phospholipase from Staphylococcus hyicus: Expression in Escherichia coli,

large-scale purification and application in esterification reactions. J. Mol.

Catalysis B: Enzymatic. 3, 13-23.

Simons, J. W., Van Kampen, M. D., Riel, S., Gotz, F., Egmond, M. R., and Verheij, H.

M. (1998). Cloning, purification and characterization of the lipase from

Staphylococcus epidermidis--comparison of the substrate selectivity with those of

other microbial lipases. Eur. J. Biochem. 253, 675-683.

52

Sinchaikul, S., Tyndall, J. D., Fothergill-Gilmore, L. A., Taylor, P., Phutrakul, S., Chen,

S. T., and Walkinshaw, M. D. (2002). Expression, purification, crystallization and

preliminary crystallographic analysis of a thermostable lipase from Bacillus

stearothermophilus P1. Acta. Crystallogr. D Biol. Crystallogr. 58, 182-185.

Sinchaikul, S., Sookkheo, B., Phutrakul, S., Pan, F. M., and Chen, S. T. (2001).

Optimization of a thermostable lipase from Bacillus stearothermophilus P1:

overexpression, purification, and characterization. Protein Expr. Purif. 22, 388-

398.

Sobek, H., and Gorisch, H. (1988). Purification and characterization of a heat-stable

esterase from the thermoacidophilic archaebacterium Sulfolobus acidocaldarius.

Biochem J. 250, 453-458.

Sommer, P., Bormann, C., and Gotz, F. (1997). Genetic and biochemical

characterization of a new extracellular lipase from Streptomyces cinnamomeus.

Appl. Environ. Microbiol. 63, 3553-3560.

Sonnet, P. E., and Gazzillo, J. A. (1991). Evaluation of lipase selectivity of hydrolysis.

J. Am. Oil Chem. Soc. 68, 11-15.

Stieb, M., and Schink, B. (1985). Anaerobic oxidation of fatty acids by Clostridium

bryantii sp. nov., a sporeforming, obligately syntrophic bacterium. Arch Microbiol.

140, 387–390.

Svensson, I., Adlercreutz, P., and Mattiasson, B. (1990). Interesterification of

phosphatidylcholine with lipases in organic media. Appl. Microbiol. Biotechnol. 33,

255-258.

53

Svetlitshnyi, V., Rainey, F., and Wiegel, J. (1996). Thermosyntropha lipolytica gen.

nov., sp. nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing

short and long chain fatty acids in syntrophic coculture with a methanogenic

archaeum. Int. J. Syst. Bacteriol. 46, 1131-1137.

Talon, R., Dublet, N., Montel, M. C., and Cantonnet M. (1995). Purification and

characterization of extracellular Staphylococcus warneri lipase. Curr. Microbiol.

30,11-16.

Tanaka, K., Yoshida, K., Sasaki, C., and Osano, Y. T. (2002). Practical asymmetric

synthesis of the herbicide (S)-indanofan via lipase-catalyzed kinetic resolution of

a diol and stereoselective acid-catalyzed hydrolysis of a chiral epoxide. J. Org.

Chem. 67, 3131–3133.

Tsuzuki W. (2005). Acidolysis between triolein and short-chain fatty acid by lipase in

organic solvents. Biosci. Biotechnol. Biochem. 69, 1256-1261.

Tyndall, J. D., Sinchaikul, S., Fothergill-Gilmore, L. A., Taylor, P., and Walkinshaw, M.

D., 2002. Crystal structure of a thermostable lipase from Bacillus

stearothermophilus P1. J. Mol. Biol. 323, 859-869.

Van Kampen, M. D., Rosenstein, R., Gotz, F., and Egmond, M. R. (2001). Cloning,

purification and characterization of Staphylococcus warneri lipase 2. Biochim.

Biophys. Acta. 1544, 229-241.

van Pouderoyen, G., Eggert, T., Jaeger, K. E., and Dijkstra, B. W. (2001). The crystal

structure of Bacillus subtilis lipase: a minimal alpha/beta hydrolase fold enzyme.

J. Mol. Biol. 309, 215-26.

54

van Tilbeurgh, H., Egloff, M. P., Martinez, C., Rugani, N., Verger, R., and Cambillau, C.

(1993). Interfacial activation of the lipase-procolipase complex by mixed micelles

revealed by X-ray crystallography. Nature. 362, 814-20.

Verger, R. (1997). Interfacial activation’ of lipases: facts and artifacts. Trends

Biotechnol. 15, 32–38.

Verger, R. and G. H. DeHaas. (1976). Interfacial Enzyme Kinetics of Lipolysis. Annu.

Rev. Biophys. Bioeng. 5:77-117.

Verger, R., Mieras, M. C., and de Haas, G. H. (1973). Action of phospholipase A at

interfaces. J. Biol. Chem. 248, 4023-4034.

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. J Mol Catal B Enzym. 9, 113–148.

Wang, N., Liu, B. K., Wu, Q., Wang, J. L., and Lin, X. F. (2005). Regioselective

enzymatic synthesis of non-steroidal anti-inflammatory drugs containing glucose

in organic media. Biotechnol. Lett. 27, 789-792.

Wei, Y., Contreras, J. A., Sheffield, P., Osterlund, T., Derewenda, U., Kneusel, R. E.,

Matern, U., Holm, C., and Derewenda, Z. S. (1999). Crystal structure of brefeldin

A esterase, a bacterial homolog of the mammalian hormone-sensitive lipase. Nat.

Struct. Biol. 6, 340-345.

Wei, Y., Schottel, J. L., Derewenda, U., Swenson, L., Patkar, S., and Derewenda, Z. S.

(1995). A novel variant of the catalytic triad in the Streptomyces scabies esterase.

Nature Struct. Biol. 2, 218–223.

55

Wei, Y., Swenson, L., Castro, C., Derewenda, U., Minor, W., Arai, H., Aoki, J., Inoue,

K., Servin-Gonzalez, L., and Derewenda, Z. S. (1998). Structure of a microbial

homologue of mammalian platelet-activating factor acetylhydrolases:

Streptomyces exfoliatus lipase at 1.9 Å resolution. Structure. 6, 511–519.

Weng, L., Hu, Z., Deng, Z., Shi, Q., Wu, S., and Li, F. (1997). Cloning and expression

in Escherichia coli of an alkaline and thermostable exolipase from Pseudomonas

pseudoalcaligenes. Wei Sheng Wu Xue Bao. 37, 434-437.

Wiegel, J. (1998). Anaerobic alkalithermophiles, a novel group of extremophiles.

Extremophiles. 2, 257-267.

Winkler, F. K., D'Arcy, A., and Hunziker, W. (1990). Structure of human pancreatic

lipase. Nature. 343, 771-774.

Wisdom, R. A., Dunnill, P., and Lilly, M. D. (1985). Enzymic interestierification of fats:

factors influencing the choice of support for immobilized lipase. Enzyme Microb.

Technol. 7, 567-572.

Woolley, P., and Peterson, S. B. (1994). “Lipases: their structure, biochemistry and

application.” Cambridge University Press. UK.

Ye P., Xu Z. K., Wu, J., Innocent, C., and Seta, P. (2006). Nanofibrous

poly(acrylonitrile-co-maleic acid) membranes functionalized with gelatin and

chitosan for lipase immobilization. Biomaterials. 27, 4169-4176.

Yemul O., and Imae T. (2005). Covalent-bonded immobilization of lipase on

poly(phenylene sulfide) dendrimers and their hydrolysis ability.

Biomacromolecules. 6, 2809-2814.

56

Yumoto, I., Hirota, K., Sogabe, Y., Nodasaka, Y., Yokota, Y., and Hoshino, T. (2003).

Psychrobacter okhotskensis sp. nov., a lipase-producing facultative psychrophile

isolated from the coast of the Okhotsk Sea. Int. J. Syst. Evol. Microbiol. 53, 1985-

1989.

Zha, D., Wilensek, S., Hermes, M., Jaeger, K-E, and Reetz, m. T. (2001). Complete

reversal of enantioselectivity of an enzyme-catalyzed reaction by directed

evolution, Chem. Commun. 24, 2664–2665.

Zhang, C., Liu, X., and Dong, X., (2004). Syntrophomonas curvata sp. nov., an

anaerobe that degrades fatty acids in co-culture with methanogens. Int. J. Syst.

Evol. Microbiol. 54, 969-973.

Zhu, B., and Panek, J. S. (2000). Total synthesis of epothilone A. Org. Lett. 2, 2575-

2578.

Zhu, B., and Panek, J. S. (2001). Methodology based on chiral silanes in the synthesis

of polypropionate-derived natural products — total synthesis of epothilone A. Eur.

J. Org. Chem. 001, 1701–1714.

57

58

Table 1.1 Changes of Gibbs free energies for several hydrogen consuming and

hydrogen releasing reactions. Table is adopted from Schink, et al. (1997).

Reaction ∆G°′

kJ/mol

Hydrogen-releasing reactions

-Primary alcohols

CH3CH2OH + H2O → CH3COO- + H+ + 2H2

-Fatty aids

+9.6

CH3CH2CH2COO- + 2H2O→ 2CH3COO- +2H+ + 2H2 +48.3

CH3CH2COO- + 2H2O→ CH3COO- + CO2 + 3H2 +76.0

CH3COO-+ H+ + 2H2O→ 2CO2 + 4H2 +94.9

CH3 CH(CH3)CH2COO- + CO2 + 2H2O→ 3CH3COO- +2H+ + H2 +25.2

Hydrogen-consuming reactions

4H2 + 2CO2→ CH3COO- + H+ + 2H2O -94.9

4H2 + CO2→ CH4 + 2H2O

H2 + S° → H2S

4H2 + SO42- + 2H+ → H2S + 4H2O

-131.0

-33.9

-151.0

Table 1.2. Families of lipolytic enzymes modified from Arpigny and Jaeger (1999).

Amino acid sequence similarities were determined with the program MEGALIGN (DNASTAR), with the first member of

each family (subfamily) arbitrary set at 100%. Abbreviations: OM, outer membrane; PHA, polyhydroxyalkanoate.

Similarity (%) Family Subfamily Enzyme-producing

strain Accession no.

Family Subfamily Properties

I 1 Pseudomonas aeruginosa*

D50587 100 True lipases

Pseudomonas fluorescens C9

AF031226 95

Vibrio cholerae X16945 57 Acinetobacter

calcoaceticus X80800 43

Pseudomonas fragi X14033 40 Pseudomonas

wisconsinensis U88907 39

Proteus vulgaris U33845 38 2 Burkholderia glumae* X70354 35 100 Chromobacterium

viscosum* Q05489 35 100

Burkholderia cepacia* M58494 33 78 Pseudomonas luteola AF050153 33 77 3 Pseudomonas

fluorescens SIK W1 D11455 14 100

Serratia marcescens D13253 15 51

59

4 Bacillus subtilis LipA M74010 16 100 Bacillus subtilis LipB C69652 17 74 Bacillus pumilus A34992 13 80 B. licheniforis U35855 13 80 5 Geobacillus

stearothermophilus L1* U78785 15 100

Geobacillus stearothermophilus P1*

AF237623 15 95

G. thermocatenulatus X95309 14 94 G. thermoleovorans AF134840 14 92 6 Propionibacterium

acnes X99255 14 100

Streptomyces cinnamoneus

U80063 14 50

7 Staphylococcus aureus M12715 14 100 S. haemolyticus AF096928 15 45 S. hyicus X02844 15 64-66# Phospholipase S. epidermidis AF090142 13 44# S. warneri AF208033 12 36 II (GDSL)

Aeromonas hydrophila P10480 100 acyltransferase

Streptomyces scabies* M57297 36 esterase Pseudomonas

aeruginosa AF005091 35 OM-bound

esterase Salmonella typhimurium AF047014 28 OM-bound

esterase Photorhabdus

luminescens X66379 28 Secreted

esterase III Streptomyces M86351 100 Extracellular

Table 1.2. Families of lipolytic enzymes (cont’d)

60

exfoliatus* lipase Streptomyces albus U03114 82 Extracellular

lipase Moraxella sp. X53053 33 Extracellular

esterase 1 IV (HSL)

Alicyclobacillus acidocaldarius

X62835 100 Esterase

Pseudomonas sp. B11-1 AF034088 54 Lipase Archaeoglobus fulgidus AE000985 48 Carboxyl

esterase Alcaligenes eutrophus L36817 40 Putative lipase Escherichia coli AE000153 36 Carboxylestera

se Moraxella sp. X53868 25 Extracellular

esterase 2 V Pseudomonas

oleovorans M58445 100 PHA-

depolymerase V Haemophilus influenzae U32704 41 Putative

esterase Moraxella sp. X53869 34 Extracellular

esterase 3 Sulfolobus

acidocaldarius AF071233 32 Esterase

Acetobacter pasteurianus

AB013096 20 Esterase

VI Synechocystis sp. D90904 100 Carboxyl esterases

Spirulina platensis S70419 50 Pseudomonas

fluorescens* S79600 24

61

62

Rickettsia prowazekii Y11778 20 Chlamydia trachomatis AE001287 16 VII Arthrobacter oxydans Q01470 100 Carbamate

hydrolase Bacillus subtilis P37967 48 p-Nitrobenzyl

esterase Streptomyces coelicolor CAA22794 45 Putative

carboxyl esterase

VIII Arthrobacter globiformis AAA99492 100 Stereoselective esterase

Streptomyces chrysomallus

CAA78842 43 Cell-bound esterase

Pseudomonas fluorescens SIK W1

AAC60471 40 Esterase III

*Lipolytic enzyme with known 3D structure, # multiple lipases similarity range

Table 1.3. Commercial lipases produced by Novo Nordisk.

Brand Name Type Of Enzyme Main Application

Lipopan® Lipase Baking industry

Lipozyme® Lipase Oils and fats industry

Novozym® 27007 Lipase Pasta/Noodles

PalataseTM Lipase Dairy industry

Clear-LensTM LIPO Lipase Personal care industry

Greasex Lipase Leather

LipolaseTM Lipase Detergent industry

LipoPrime® Lipase Detergent industry

NovoCorTM AD Lipase Leather industry

Novozym® 735 Lipase Textile industry

Novozym® 871 Lipase Pet Food Industry

63

64

Table 1.4. The most thermophilic lipases.

Source Toptima pHoptima Stability Ref.

Thermosyntropha

lipolytica

96ºC 9.0-9.6 LipA: t1/2 6 h at

100°C, 24h at 75°C.

LipB: t1/2 2h at 100°C,

24h at 75°C at pH 8

This study

Burkholderia

cepacia

90ºC 11 t1/2 13 h at 90ºC Rathi, 2000

Geobacillus

thermoleovorans

75°C

8.0

t1/2 =1 hr at 60°C at

pH 7.5

Cho, 2000

Bacillus

sp. THLO27

68-

70°C

7.0

t1/2=2 hr at 60°C &

75°C at pH 8.0

Dharmsthiti

and Luchai.

1999.

Geobacillus

thermocatenulatus 60°C

-70°C

BTL2

8-9 t1/2= 30 min at 60°C

at pH 9.0

Rua Luisa,

1997

Geobacillus

stearothermophilus

L1

60-

65°C

9-10 retains 30% activity

after 30min at 65°C

at pH 8.0

Kim, 1998

Myroides

odoratimimus (bas

Flavobacterium

odoratum)

60°C 10.0 T1/2=10.2 min at 70°C

at pH 8.5

Labuschagne,

1997

Bacillus sp. J33 60°C 8.0 T1/2=12 h at 60ºC Nawani and

Kaur. 1999.

Bacillus sp. TG4 60°C 9.0 Retains 80% activity

after 30 min at 60°C

Bell, 1999

Geobacillus

stearothermophilus

P1

55°C 8.5 T1/2=2 h at 65°C at

pH 8.5

Sinchaikul,

2001

Table 1. 5. Properties of bacterial lipases from Gram positive and Pseudomonas sp.

Origin pH*

T (°C) Size

(kDa)

Cloning Host

/Vector

Reference

Geobacillus

thermoleovorans

8.0 75 46 E. coli JM109

PUC19

Cho, 2000

Bacillus sp. THLO27 7.0 68-70 69 nd Dharmsthiti, 1999

Geobacillus

thermocatenulatus

8.0 60 16,

40

E. coli BL321

pCYTEXP1

Rua Luisa,1997

Schmidt-D, 1994

Geobacillus

stearothermophilus L1

9.0 60-65 43 E. coli RR1

PUC19

Kim, 1998

Bacillus sp. J33 60 45 nd Nawani, 1999

Bacillus sp. TG4 9.0 60 E. coli DH5α

λzap11

Bell, 1999

Geobacillus

sterothermophilus P1

8.5 55 43 E. coli M15

PQE60

Sinchaikul, 2001

& 2002

Bacillus sp. H257 7.0 37 27.4 E. coli DH1 Kitaura, 2001

65

PACYC184

B. subtilis 168 10.0 35°C 19 E. coli/pLIP1 Dartois, 1994

Lesuisse, 1993

Staphylococcus

aureus

6 37°C 34,

46

E. coliDH5α Simons, 1996

Gotz, 1998

S. simulans 8.5 37°C 160 Nd Sayari, 2001

S. hyicus

8.5 46 E. coli DH5α

PBR322

Gotz, 1998

Simmons, 1997

S. epidermidis 6 37°C 43 E. coli DH5α

PET15b

Simmons, 1998

S. haemolyticus 8.5 28°C 45 E. coli XL1

pBluescriptII

Oh, 1999

S. wareni 9.0 25°C 45 E. coli DH5α

pRB473

Van kampen,2001

Talon, 1995

66

67

Pseudomonas

sp. ATCC 21808

Stable

5-10

Stable

50-60°C

35 Kordel, 1991

Pseudomonas

fluorescence

8-10 55°C 33 nd Kojima, 1994

Pseudomonas sp. 7.0 45-60°C 40 Dong, 1999

P. aeruginosa TE3285 7-9 45-50°C 30 E. coli 1100

PUC18

Chihara-, 1992

P. fluorescens HU380 8.5 45°C 64 E. coli JM109

pBluescriptII

Kojima, 2003

Pseudomonas

pseudoalcaligens

6-10 40°C 32 E. coli HB101

PUC118

Lin, 1996

Weng, 1997

P. aeruginosa YS-7 7 20-55°C 40 Shabtai, 1992

Pseudomonas sp.B11-1 8 37°C 33.7 E. coli C600

PUC118

Choo, 1998.

Figure 1.1.

The lipase reaction catalyzing the hydrolysis and synthesis of lipids. The

hydrolysis of lipids into glycerol and fatty acids occurs in aqueous solution. The reverse

synthesis reaction occurs in water restricted organic solvent.

68

3H2O

R1

R2

R3

+

H

H

H

H H R1OH

Lipase + H R2OH

OH H R3

H Triacylglycerol Glycerol Fatty acids

69

Figure 1.2.

The effect of substrate concentration on hydrolysis rate. Thermosyntropha.

lipolytica lipase; LipA (A) (this study) and Alicyclobacillus acidocaldarius esterase;

EST2 (B) (adopted from Chahinian, et al., 2005). The Activity is expressed as

percentage of maximal activity (vmax).

70

0

20

40

60

80

100

0 1 2 3 4

0

20

40

60

80

100

0 0.5 1

A

p-nitrophenyl laurate (mM) v

p-nitrophenyl butyrate (mM)

B

71

Figure 1.3.

(A) Model for describing lipase kinetics acting on insoluble substrate at the

interface. E is the enzyme in closed conformation, E* is the activated enzyme in open

conformation (penetrated enzyme concentration), E*S activated (penetrated) enzyme

substrate complex, P is final product k1 and k-1 are the rate constants of forward and

reverse reactions, respectively, kcat is the catalytic constant, on this case it is the same

as k2, kp and kd are penetration and desorption rate constants, respectively Figure is

adopted from Verger, et al. (1973). (B) Kinetic mechanism (Ping Pong Bi Bi) of

lipase-catalyzed reactions involving multiple substrates/products. Using Cleland’s

nomenclature (E: enzyme; Es: ester moiety; Al: alcohol moiety; Ac: acid moiety; W:

water; F: acyl enzyme complex; i = 1, 2, … , I; j = 1, 2, … , J); Aci denotes an acid

moiety of the i-th type, Alj an alcohol moiety of the j-th type, Esi,j an ester moiety

containing an acid residue of the i-th type and an alcohol residue of the j-th type. Figure

is adopted from Malcata, et al. (2000).

72

E

kp kd

E*

S

E*S

P

k1

K-1 kcat

A

Water-lipid interface

Aqueous solution

B

73

Figure. 1.4.

(A) The common lipase fold. Arrows indicate β strands and rectangles indicate α

helices. β strands are numbered according to the nomenclature of the α/β hydrolase fold.

Secondary structural elements shown in black or white (strands β3-β7 and helices B and

C) occur in all lipases; those shown in gray (strand β2 and helices A, D, and F) occur in

most. Helices A and F are on the concave side of the β sheet; the other helices are on

the convex side. Helix D is often composed of only one (distorted) turn. Figure is

adopted from Schrag and Cygler (1997). (B) Schematic diagram for serine

hydrolase’s reaction mechanism catalyzing esterification or hydrolysis of fatty

acid alcohol. The tetrahedral intermediate stabilized by hydrogen bonding to form the

oxyanion hole is shown. Figure is adopted from Raza, et al. (2001).

74

A

N

A B C

β2 β3 β4 β5

75

C

S H

D

D

F

β6 β7

B

HN

HN

76

Figure 1.5.

Structure of P. aeruginosa lipase. (A) Schematic view of the secondary structure

elements. The ribbon representation was made using MOLSCRIPT (Lang, et al., 1996);

α-helices, β-strands, and coils are represented by helical ribbons, arrows, and ropes,

respectively. α-Helices belonging to the cap domain involved in substrate binding are

shown in red. The position of the -helical lid is highlighted with the label LID. The

phosphonate inhibitor covalently bound to the nucleophile Ser82, the calcium ion, and

the disulfide bridge are in ball and stick representation in cyan, black, and yellow,

respectively. (B) Secondary structure topology diagram. The catalytic triad residues

(Ser82, Asp229, and His251) and the position of the disulfide bridge are indicated, and a

comparison with the canonical α/β hydrolase fold is given. α-Helices and β-strands are

represented by rectangles and arrows, respectively. G1 and G2 are 310 helices and are

represented by squares. Locations where insertions in the canonical fold may occur are

indicated by dashed lines. Figures are adopted from Nardini, et al. (2000).

77

78

Figure 1.6.

The secondary structure topology of G. stearothermophilus P1 lipase is

compared to that of the canonical α/β hydrolase fold. (A) Secondary structure

topology, showing the general α/β hydrolase fold including the catalytic triad and the

zinc-binding structural elements and residues indicated in black and red, respectively.

Alpha helices and beta strands are represented by rectangles and arrows, respectively.

New structural elements, strand b1, helix α3 and strand b2 are shown in red. BSP lacks

the N-terminal antiparallel beta sheet of the canonical fold. Helix α6 and the adjacent

loop region make up the lid which, in its closed position, isolates the substrate-binding

cleft from solvent. (B) Secondary structure topology diagram of the canonical α/β

hydrolase fold. Broken lines indicate possible sites of insertions. The heavy line

depicts the position of the new deviation from the known fold. Figures are adopted from

Tyndall, et al. (2002).

79

A

B

N

80

Figure 1.7.

Ribbon diagram of both lipases BSP and BSL from G. stearothermophilus. (A)

Structure of G. stearothermophilus P1 lipase. Alpha helices, beta strands and loops

are shown in blue, red and yellow, respectively. The labelled residues, from left to right,

are the calcium-binding residues, Glu360, Gly286, Pro366 and Asp365 with calcium in

green; the catalytic triad, Asp317, His358 and Ser311, and the zinc-binding residues,

Asp61, His87, Asp238 and His81 with zinc in orange adopted from (Tyndall, et al.,

2002). Structure of G. stearothermophilus L1 lipase. Secondary structural elements

are labeled on the drawing. Calcium and zinc atoms are represented as cyan and

purple balls, respectively. Side chains of catalytic triad residues (Ser-113, His-358, and

Asp-317) are shown in a ball-and-stick representation. Figures are adopted from Jeong,

et al. (2002).

81

A

B

N

82

CHAPTER 2

PURIFICATION AND CHARACTERIZATION OF TWO HIGHLY THERMOPHILIC

ALKALINE LIPASES FROM THERMOSYNTROPHA LIPOLYTICA1

1Salameh, M. and J. Wiegel. 2006. To be submitted to Applied and Environmental Microbiology.

83

ABSTRACT

Thermosyntropha lipolytica DSM 11003 is an anaerobic thermophilic alkalitolerant

bacterium which grows syntrophically with methanogens on lipids, utilizing the fatty acid

moieties but not the glycerol. Two lipases, LipA and LipB, were purified from culture

supernatants to gel electrophoretic homogenety by ammonium sulfate precipitation and

hydrophobic interaction column chromatography. The apparent molecular weights of

LipA and LipB determined by SDS-PAGE were 50 and 57 kDa, respectively. The

temperature optima of LipA and LipB were each near 96°C, which is the highest so far

known among lipases. The pH25°C optima of LipA and LipB were 9.4 and 9.6,

respectively. They were also very thermostable, and LipA and LipB retained 50%

activity after 6 and 2 hours incubation at 100 °C, respectively. In general, both enzymes

preferred glycerides with long chain fatty acids, with maximum activity exhibited toward

trioleate (C18:1). Among the p-nitrophynyl (pNP) esters tested, pNP laurate exhibited the

highest activity. Thin layer chromatography results showed that both lipases catalyze

the hydrolysis of ester bonds at position 1 and 3. The activity of both lipases was totally

inhibited by 10mM PMSF and 10 mM EDTA. Metal analysis showed that LipA and LipB

each contain one atom of Ca2+ and one of Mn2+. The addition of 1 mM MnCl2 to dialyzed

enzyme preparations enhanced the activity of both lipases by three fold and increased

their thermal stability by 4 h to 34 h at 60 °C

84

INTRODUCTION

Thermosyntropha lipolytica is an anaerobic, thermophilic, organotrophic, lipolytic,

and alkalitolerant gram type positive bacterium, and was isolated from an alkaline hot

springs of Lake Bogoria, Kenya, using minimal media plus olive oil (52). The bacterium

was isolated with the specific intent of finding bacterial lipases exhibiting high stability

and activity at high temperatures and high pH values. True lipases (carboxyl ester

hydrolases, E.C. 3.1.1.3) are enzymes that generally catalyze the synthesis and

hydrolysis of long chain fatty acid esters (14, 44). They are ubiquitous in nature,

produced by animals, plants, fungi, as well as bacteria but (it is assumed that the

authors have access to all public information!) have not yet been reported in archaea.

Lipases are activated at the water-lipid interface, they show little activity when the

substrate is in the monomeric form and the activity increases dramatically above the

solubility limit where the substrate start to form emulsions. This fact has led to the

emergence of a phenomenon known as interfacial activation, which describes substrate

emulsions as a necessity for maximum lipolytic activity (8, 14, 44, 55) for the majority of

lipases. However, there are a few exceptions that are not interfacially activated despite

homology to other known lipases (19, 31, 33).

All lipases have a conserved 3-D structure (2), share the α/β hydrolase fold, and

have the same catalytic mechanism. The catalytic center contains the catalytic triad

serine-histidine-aspartic or glutamic acid. On the other hand, lipases can have little if

any similarity in their primary sequence, molecular mass, pH and temperature optima,

substrate and positional specificity, cofactors and cellular location (2, 17).

85

Lipases are among the most versatile and most studied of all enzyme classes.

They are used in a number of applications in various industries including

pharmaceutical, dairy, detergent, cosmetic, oleochemical, fat processing, leather, textile,

cosmetic and paper industries (4, 7, 21, 22, 45, 56).

Many enzymes have been found to be fairly thermo-and-alkali stable,

including those from mesophilic organisms. However, It is likely that enzymes produced

by thermophilic, alkalitolerant bacterium will be more thermostable at alkaline pH while

exhibiting high specific activities at elevated temperature (>70 °C) and alkaline pH (> 9).

While a few lipases have been reported from aerobic thermoalkaliphilic bacteria (5, 26,

27, 30, 41), here we report on the first characterization and purification of lipases from

an anaerobic thermophilic bacterium, Thermosyntropha lipolytica.

MATERIALS AND METHODS

Culture and growth conditions. Thermosyntropha lipolytica DSM 11003T was grown

in a basal medium containing 0.75% yeast extract as carbon and energy sources under

nitrogen gas phase. The basal medium contained (per liter) 0.3 g of K2HPO4, 0.3 g of

KCl, 0.5 g of NaCl, 1.0 g of NH4Cl, 0.1g of MgCl2·6H2O, 0.02 g of CaCl2·2H2O, 3.0 g of

NaHCO3, 3.0 of Na2CO3, 0.5 g of Na2S·9H2O, 0.15 g of cysteine, 2 ml of vitamin

solution and 2.5 ml of trace element solution (15). The pH25°C of the medium was

adjusted at 8.2 and the growth temperature was 60 °C. To obtain biomass for lipase-

related measurements, 500 ml of anaerobic medium was inoculated with 5% (v/v) of a

twelve hour old culture of T. lipolytica, the pH of the medium was adjusted by using 0.2

M HCl and 0.2 N NaOH and incubated at 60 °C. Every time a sample was withdrawn,

86

the pH of the culture was adjusted. After measuring the O.D., the sample was

centrifuged and then prepared for protein quantification and lipase assay.

Lipase assay. Lipase was assayed spectrophotometrically using p-nitrophenyl laurate

(pNPL) (Sigma) and p-nitrophenyl palmitate (pNPP) (Sigma) as substrates (58). The

reaction mixture contained 100 µl of enzyme solution, 25 µM MnCl2, 1.088 ml of freshly

prepared buffer containing 100 mM HEPES, 100 mM TAPS and 100 mM CAPS buffer

(Sigma), and 12 µl of 300 mM pNPL or pNPP in acetonitrile (final concentration in the

assay, 3 mM). The assay was typically carried out for 20 minutes at 96 °C (the assay

was linear for upto 45 min). The reaction mixture was then cleared by centrifugation,

and the A405 of the liberated p-nitrophenol was determined. One unit is defined as the

amount of the enzyme catalyzing the release of 1 µmol of p-nitrophenol per min (ε=1.82

× 104 M-1 cm-1) from pNP-laurate/palmitate.

Purification of lipases. All purification steps were performed at room temperature.

Medium, 20 l, was inoculated with 2 l of exponentially growing pre-culture. The pH was

adjusted to 8.225°C (7.660°C), and the growth temperature was 60 °C. After 18 h, the cells

were separated by Amicon hollow fiber filter with a one million Da cut off, and the

supernatant was recovered. The extracellular lipase was then concentrated through

filtration using a 10 kDa Amicon hollow fiber filter (Millipore), and precipitated using

stepwise saturation to 60% and 75% ammonium sulfate. The precipitate was collected

by centrifugation and dissolved in 10 mM sodium phosphate buffer, pH 8.0, and applied

onto a 30 ml Octyl Sepharose fast flow column (Amersham Biosciences). The column

was pre-equilibrated with 20 mM Tris buffer, pH 8.0, containing 2 M (NH4)2SO4 (buffer

A). The bound protein was eluted with a decreasing gradient from 450 ml of buffer A to

87

450 ml of 20 mM Tris buffer, pH 8.0. Fractions containing high specific lipase activity

(1.8 M and 0.5 M (NH4)2SO4) were pooled, desalted, and concentrated using a filter

membrane (Millipore). The lipases were tested for purity on native and SDS poly

acrylamide gel electrophoresis as described by Sambrook and Ausubel (3, 43). Gels

were stained with GelCode® Blue Stain Reagent (Pierce). The molecular mass of the

enzyme was estimated by interpolation graphically from a graph log of molecular mass

versus relative migrations (RF values).

Protein concentration. Protein concentrations were determined with the bicinchoninic

acid (BCA) protein assay kit (Pierce) following the manufacturer instructions.

The effect of temperature on activity and stability. A temperature gradient incubator

(Scientific Industries Inc., Bohemia, NY, USA) was used to determine the temperature

optima of both lipases. The enzyme assay was performed at pH25°C 9.4 and 9.6, which

were equal to pH80°C 8.5 and 8.8 for LipA and LipB, respectively (57). The enzyme

assays were performed as described above, using mixed buffer of 100 mM HEPES, 100

mM TAPS and 100 mM CAPS.

Thermostability was analyzed by measuring the residual activity after incubating

the enzyme in Tris buffer pH25°C 8.0 at 60, 75, 100°C for various times in series of

sealed 2 ml serum bottles, from which 3 bottles were sacrificed for every time point.

Each sample contained 15 µg of protein in 0.6 ml of 100 mM Tris buffer, pH 8.0. After

incubation, the samples were concentrated and desalted when necessary by using 10

kDa cut off centricon filter tubes (Millipore) they were then assayed in triplicate for lipase

activity, and the protein was quantified.

88

The effect of pH on activity and stability. To determine the pH optima of both

enzymes, standard lipase assays were carried out at a pH25°C range of 3.0 to 11.5. A

mixture of the following buffers was used, each at 100 mM concentration: glycine–HCl,

sodium acetate, MES, HEPES, TAPS and CAPS. Because the substrate pNPL was not

stable at pH25°C over 10, triolein was used and the liberated fatty acids were measured

by the NEFA C kit (Waco USA).

For stability measurements, 15 µg of purified LipA and LipB were stored in 0.6 ml

aliquots of different buffers ranging from pH 2.0 to pH 12. After 36 hours of incubation

at room temperature, the samples were concentrated and assayed for lipase activity;

the assays were in triplicate.

Determination of substrate specificity. Substrate specificity and chain length

selectivity were determined spectrophotometerically using a variety of p-nitrophenyl

esters (Sigma) ( p-nitrophenyl acetate, p-nitrophenyl butyrate, p-nitrophenyl laurate, p-

nitrophenyl myristate, p-nitrophenyl palmitate). Triglycerides (Sigma), including tributyrin

(C4), tricaproin (C6), tricaprylin (C8), tricaprin (C10), trilaurin (C12), trimyristin (C14),

tripalmitin (C16), tristearin (C18) and triolein (C18:1) was determined by the NEFA C kit.

Long chain glycerides were dissolved in acetonitrile. The assay contained 5 mM final

concentration of substrate and 1mM MnCl2.

Determination of positional specificity. The positional specificity of LipA and LipB

was analyzed by thin layer chromatography (TLC) using a modified version of

procedure reported by Lesuisse et al. (31). The enzymes assays were carried out at the

temperature and pH optima of both lipases for 8 hours. The assay solution contained 5

mM triolein and 1 mM MnCl2 in a 100 mM of TAPS buffer. The assay mixture was

89

sonicated for 30 sec before the addition of 50 µg of the purified enzyme, and was

terminated by rapid cooling in an ice bath. The reaction products were extracted with 2

ml of cold diethylether. The extract was concentrated by evaporation and applied on

silica gel plate (Fisher Scientific). Monolein, 1,3 diolein, 1,2 diolein and triolein (Sigma)

were used as standard reference. The plates were developed with a mixture of

chloroform:acetone:acetic acid (96:4:1). The separated esters were visualized by

spraying the plates with 0.1% iodine in chloroform.

The effect of metal ions, EDTA and phenylmethanesulfonoylfluoride (PMSF). To

determine the effect of metal ions, EDTA and PMSF on activity, different concentrations

up to 10 mM were added directly to the assay solution. The standard lipase assay was

followed as described earlier. In addition to the standard assay, the effect of calcium,

manganese and iron metals along with EDTA and PMSF were tested with triolein as

substrate and the free fatty acids quantified using the NEFA C kit .

RESULTS

Lipase level in the culture.

It was already known that lipase was formed constitutively by this organism (52).

However, we showed here the maximum formation of lipase at different growth pH (Fig.

2.1). At pH60ºC 7.6, which is the optimum pH for growing T. lipolytica, the maximum

specific activity using pNPL was 0.15 U.mg-1 after 15 to18 hours of growth. At pH60ºC 9.0,

the maximum specific activity was 0.12 U.mg-1, and it was achieved after 21 hours of

growth.

90

Purification of lipases LipA and LipB.

The lipases were isolated from the culture supernatant using ammonium sulfate

precipitation and hydrophobic interaction chromatography. A purification of 140 fold was

achieved (Table 2.1). Two different lipases were eluted from the Octyl Sepharose

column. Lipase A (LipA) was weakly bound to the column and was eluted at 1.8 M

ammonium sulfate. The other lipase (LipB) was more strongly bound and eluted with 0.5

M ammonium sulfate. LipA and LipB have molecular masses of 50 and 57 kDa,

respectively, as determined by SDS-PAGE (Fig. 2.2). Their N-terminal sequences were

NGGGATLPLQTSGVLTAGFAP and VKVMATLPADYVAQVIENVKR, respectively,

suggesting that they are two distinct enzymes.

LipA and LipB were found to be cold labile, irreversibly inactivated when frozen.

A 75% and 90% catalytic activity of LipA and LipB were preserved, respectively, when

frozen in solution that contains 40% glycerol (v/v) and 2 mg/ml bovine serum albumin.

Effect of pH and temperature on enzymes activity and stability.

The maximal activity of LipA and LipB in 20 min assay was at 96 °C (Fig. 2.3) although

the temperature optima of both enzymes vary according to the assay buffer. Using

TAPS buffer instead of the combination buffer at pH 9.0, the temperature optima of LipA

shifted to 98 ºC, while LipB temperature optimum shifted down to 90 ºC. The pH25°C

optima of LipA and LipB were 9.4 and 9.6, respectively (Fig. 2.4). The buffer used also

affected the measured pH optima. Using TAPS buffer, the temperature optima was in

the range of 8.8 to 9.1 for both enzymes. Because p-nitrophenyl laurate is unstable at

temperatures above 98 °C and above pH 10.0, the true substrate trioleate was used at

91

high temperature and pH values. The liberated fatty acids were quantified by using

NEFA C kit.

LipA and LipB showed maximum thermostability in the presence of 0.5 M ammonium

sulfate. They retained 50% activity after incubation at 100 °C for 6 and 2 hours,

respectively, and 48 and 72 hours at 75 °C and 60 °C, respectively (Fig. 2.5). Both

enzymes were unstable when incubated at acidic pH (below 7.0) (Fig. 2.5), but both

were very stable at alkaline pH, retaining 100% of their activity after incubation for 72

hours at pH 7.0 to pH 12.0.

Substrate specificity and positional specificity.

The two enzymes were assayed using triglycerides and synthetic nitrophenyl substrates

(Fig. 2.6). Both showed high activity toward substrates with long chain fatty acids (C12-

C18) and showed higher activity toward substrates with unsaturated fatty acids as in

triolein, both enzymes exhibited low activity toward soluble substrates of short chain

fatty acids (≤ C6), which is consistent with the definition of a true lipase. The hydrolytic

products of triolein after 2, 4 and 8 hours assays appeared to be 2-monoolein, indicating

a preference for digestion of the sn-1,3-position of triolein (Fig. 2.7).

The effect of metal ions, PMSF and EDTA on enzymes activity.

The catalytic activity of LipA and LipB of T. lipolytica were neither enhanced nor

inhibited by the presence of CaCl2 (1mM to 10 mM, Table 2.2). However, a 15%

decrease in activity was noticed when 25 mM of CaCl2 was added to the assay mixture.

LipA and LipB showed various degrees of inhibition by all metal ions tested (Table 2.2),

and the inhibition was greater as the concentration of the ion increased. Manganese (in

the form of MnCl2) at concentrations between 0.1 to 2 mM increased the activity of LipA

92

and LipB by 3-fold compared to assays conducted in the absence of metal ions or in the

presence of calcium (0.1-10 mM), which had no effect. The effect of manganese was

confirmed by carrying out lipase assays using triolien as substrate and quantifying the

liberated fatty acids. In the presence of 5 mM EDTA, nearly half the activity of LipA and

LipB was lost, and the activity was completely diminished in the presence of 10 mM

EDTA. These results indicate that LipA and LipB might require a certain metal for

activity.

To determine whether LipA and LipB might contain Mn-binding sites similar to

Zn-binding sites found in thermophilic lipases from G. stearothermophilus, we

performed metal analyses (Inductively Coupled Plasma-Emission Spectrometry) using

two enzyme concentrations after dialysis against distilled water. The values corrected

against the values of the dialysis buffer yielded 0.92 ± 0.1 mol Ca2+ and 0.6 ± 0.06 mol

Mn2+ ions per mol of enzyme. Since the addition of manganese ions resulted in a three

fold activation of the enzyme, it is assumed that the native enzyme contains 1 mol Mn2+

ion per mol lipase, and that some is lost during the purification procedure.

Lipases are members of the serine hydrolases, where serine is an essential

residue for their catalytic activity. We tested the sensitivity of LipA and LipB to different

concentrations of the serine inhibitor, phenylmethanesulfonoylfluoride (PMSF). The

activity of both enzymes was inhibited at 10 mM PMSF concentration (Table 2.2).

DISCUSSION

Two highly thermostable enzymes present: T. lipolytica produced two extracellular

lipases. These appear to be encoded by two distinct genes, as indicated by the

different biochemical properties and different N-terminal sequences. Maximal activity

93

was observed at 96 °C (in a 20 min assay) for both enzymes but the precise optima

depended on the assay buffer used. A temperature maximum of 96 °C (while using the

true substrate triolein instead of the usually used p-nitrophenyl) indicates that these two

lipases are the most thermophilic ones so far reported. The only other lipase with such

high thermal activity and stability was surprisingly found in the mesophilic Burkholderia

cepacia (39). However, comparing the physical properties of these lipases is quite

difficult due to the differences in the assay substrates and procedure. Very few

thermophilic lipases have been characterized and none of them exhibit the high

thermostability of LipA and Lipb (26, 36, 49). The only lipase that has a similar

thermostability to LipA and LipB is the lipase from Burkholderia cepacia, it retained 50%

activity after 13 hours incubation at 90 ºC.

Role of metal-content on activity and stability of LipA and LipB: Many bacterial

lipases for which structures are available contain a Ca2+-binding site. These include the

enzymes from Chromobacterium viscosum (28), P. aeruginosa (35), and Burkholderia

(bas Pseudomonas) glumae (BGL) (37)). Many lipases have been shown found to

require certain metal for activity and/or to enhance activity and (thermo)stability. The

two thermophilic lipases from Geobacillus (bas Bacillus) stearothermophilus P1 (54) and

L1 (24) contain zinc as well as calcium binding sites, which is unique among the lipases

that have been characterized so far. Zinc is believed to enhance the thermostability of

these enzymes. In addition the activity of G. thermoleovorans ID-1 thermophilic lipase

(30) was reported to be enhanced by calcium and zinc ions. The activity of other lipases

from Pseudomonas sp. (13, 38), Acinetobacter sp. (50) B. licheniformis (25), and B.

subtilis 168 (31) were also found to be enhanced by calcium. Staphylococcal lipases

94

were found to require calcium for activity and stability (40, 16). Kim et al. showed by

means of fluorescence emission kinetics that calcium enhanced the thermostability of G.

stearothermophilus lipase (26).

The importance of calcium ions and its involvement in stabilizing the tertiary

structure of the enzyme was observed in the crystal structure of B. glumae lipase (37). It

was observed that a calcium ion forms ligands with a number of adjacent residues at

the active site. Loss of the calcium ion, through either pH change or mutation to a

residue that affects the calcium interactions has been proposed to disrupt the enzyme

structure and decrease its thermal stability as observed in Staphylococcus hyicus (48).

However, calcium neither enhanced nor reduced the activity and/or stability of LipA and

LipB. When incubating the pure enzymes without any salt or metal ion additions, LipA

and LipB both retained 50% activity after 1, 20, and 30 hours incubation at 100 °C, 75

°C and 60 °C, respectively. In contrast, manganese ions specifically enhanced the

activity of both lipases by 3-fold (Table 2.2). Metal analysis confirmed that both lipases

bind one manganese ion but that it might not be tightly bound. Calcium ion, on the other

hand, is more tightly bound. Moreover, the effect of manganese on thermostability of

LipA and LipB i.e., manganese extended the half life of both enzymes by 4 hours at

60°C and 75 °C, suggest that LipA and LipB contain a manganese-coordinated domain

similar to the Zn-coordinated domain found in G. stearothermophilus lipases. The

crystal structure of G. stearothermophilus lipase indicated that it contains a zinc-

coordinated extra domain which makes tight interactions with the loop extended from the

C terminus of the lid helix and makes strong hydrophobic interactions with its

neighboring domains including the core domain. It has been proposed that these

95

interactions lead to rigid packing of the active site and appeared to play important role in

the enzyme catalytic activity and thermostability of the enzyme; this might explain the

optimum activity at 65 °C. The lid apparently requires high temperatures in order to

move away and to expose the active site (24). Manganese in LipA and LipB might have

the same effect as zinc in contributing to the enzyme activity and stability at very high

temperatures providing tight interactions with the core domain, as well as with the

helices and loops around the lid. Other structural features which were observed and

studied in thermophilic proteins that could contribute to the thermostability of LipA and

LipB have not been investigated in detail due to lack of gene sequence, such as high

hydrophobic interactions, higher hydrogen bonding, amino acids substitution and high

salt bridges content among others (10, 11, 18, 23, 32, 42, 53).

The most profound effect on thermostability was noticed with the addition of 0.5

to 2 M of ammonium sulfate to the enzyme solution prior to assaying activity. The half-

lives of LipA and LipB were extended at all temperatures, and most noticeably was 18

hours increase at 60 °C. Benjwal and coworkers showed that molar concentrations of

NaCl or Na2SO4 increase the apparent melting temperature of a lipoprotein up to 20 °C

and that these salts decelerated protein unfolding (6). Similar effects have been also

reported for other enzymes (1, 20, 51). Ammonium sulfate is a chaotropic agent. It

increases the chaos (entropy) in water, and thereby increases hydrophobic interactions,

the proteins become less flexible, which stabilizes the tertiary structure and lowers

domain unfolding.

Substrate specificity and true lipases: LipA and LipB had a similar preference for

long chain fatty acids (LCFA) esters and especially toward unsaturated ones present in

96

triolien (C18). The preference toward LCFA was also observed with p-nitrophenyl esters

and showed maximum activity toward p-nitrophenyl laurate (C12). It is important to

emphasize that the substrate specificity preference of LipA and LipB is different from all

other described thermophilic lipases. For example, the thermophilic lipase from the

aerobic G. thermoleovorans ID-1 showed maximum activity toward tricaproin (C6) and p-

nitrophenyl caproate (C6) (9). G. stearothermophilus P1 lipase showed maximum

catalytic activity toward p-nitrophenyl caprate (C10) and tricaprylin (C8) (49). The G.

stearothermophilus L1 lipase is most active with p-nitrophenyl caprylate (C8) and

tripropionin (C3) (27), indicating that it may be an esterase and not a true lipase.

Similarly, the G. thermocatenulatus enzyme showed maximum activity toward p-

nitrophenyl butyrate and tributyrin (C4) (47).

LipA and LipB are lipases that prefer to hydrolyse ester bonds at position 1 and 3

of triglycerides. This specificity is similar to that exhibited by other lipases such as those

from Bacillus sp. THL027 (12), G. thermocatenulatus (46), B. subtilis 168 (31), and from

some Gram-negative bacteria (29, 34).

Most of the lipases that are utilized commercially are from Pseudomonas species,

presumably because these lipases were the first to be isolated, cloned, and

characterized. Their 3-D structures were also among the earliest to be revealed.

Moreover, most of these lipases have undergone a variety of molecular modifications to

produce effective and efficient enzymes with more desirable characteristics (17, 22).

However, the properties of LipA and LipB should make these enzymes of great interest

for specific industrial applications. Their properties extend the diversity of lipases and

97

indicate that extremophilic bacteria are a rich source of biotechnologically-interesting

enzyme, and particularly lipases.

98

REFERENCES

1. Arakawa, T., and M. Tokunaga. 2004. Electrostatic and hydrophobic interactions

play a major role in the stability and refolding of halophilic proteins. Protein Pept.

Lett. 11:125-132.

2. Arpigny, J. L., and K-E. Jaeger. 1999. Bacterial lipolytic enzymes: classification

and properties. Biochem. 343:177–183.

3. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. A. Smith, J. Seidman

and K. Struhl. 1995. In Current Protocols in Molecular Biology, Greene Publ., New

York.

4. Balcão, V., M. Paiva, and F. X. Malcata. 1996. Bioreactors with immobilized

lipases: State of the art. Enzyme Microb. Technol. 18:392–416.

5. Bell, P. J. L., N. Nevalainen, H. W. Morgan, and P. L. Bergquist. 1999. Rapid

cloning of thermoalkalophilic lipases from Bacillus sp. Using PCR. Biotech. Lett.

21:1003-1006.

6. Benjwal, S., S. Jayaraman, and O. Gursky. 2005. Electrostatic effects on the

stability of discoidal high-density lipoproteins. Biochemistry. 44:10218-26.

7. Bornscheuer, U. T., C. Bessler, R. Srinivas, and S. H. Krishna. 2002.

Optimizing lipases and related enzymes for efficient application. Trends Biotechnol.

20:433-437.

8. Brzozowski, A. M., U. Derewenda, Z. S. Derewenda, G. G. Dodson, D. M. -

Lawson, J. P. Turkenburg, F. Bjorkling, B. Huge-Jensen, S. A. Patkar, and L.

Thim. 1991. A model for interfacial activation in lipases from the structure of a

fungal lipase-inhibitor complex. Nature. 351:491 494.

99

9. Cho, A. R., S. K. Yoo, and E. J. Kim. 2000. Cloning, sequencing and expression

in Escherichia coli of a thermophilic lipase from Bacillus thermoleovorans ID-1.

FEMS Microbiol. Lett. 186:235-238.

10. Criswell, A. R., E. Bae, B. Stec, J. Konisky, and G. N. Phillips. 2003. Structures

of thermophilic and mesophilic adenylate kinases from the genus Methanococcus.

J. Mol. Biol. 330:1087-99.

11. Dello Russo, A., R. Rullo, G. Nitti, M. Masullo, and V. Bocchini. 1997. Iron

superoxide dismutase from the archaeon Sulfolobus solfataricus: average

hydrophobicity and amino acid weight are involved in the adaptation of proteins to

extreme environments. Biochim. Biophys. Acta. 1343:23-30.

12. Dharmsthiti, S., and S. Luchai. 1999. Production, purification and

characterization of thermophilic lipase from Bacillus sp. THL027. FEMS Microbiol.

Lett. 179:241-246.

13. Dong, H., S. Gao, S. Han, and S. Cao. 1999. Purification and characterization of a

Pseudomonas sp. lipase and its properties in non-aqueous media. Biotechnol.

Appl. Biochem. 30:251-256.

14. Ferrato, F., F. Carriere, L. Sarda, and R. Verger. 1997 A critical reevaluation of

the phenomenon of interfacial activation. Methods Enzymol. 286: 327-347.

15. Freier, D., C. P. Mothershed, and J. Wiegel. 1988. Characterization of

Clostridium thermocellum JW-20. Appl. Environ. Microbiol. 54:204-211.

16. Gotz, F., H. M. Verheij, and R. Rosenstein. 1998. Staphylococcal lipases:

molecular characterization, secretion, and processing. Chem. Phys. Lipids. 93:15-

25.

100

17. Gupta, R., N. Gupta, and P. Rathi. 2004. Bacterial lipases: an overview of

production, purification and biochemical properties. Appl. Microbiol. Biotechnol.

64:763-781.

18. Haney, P., J. Konisky, K. K. Koretke, Z. Luthey-Schulten, and P.G. Wolynes.

1997. Structural basis for thermostability and identification of potential active site

residues for adenylate kinases from the archael genus Methanococcus. Proteins:

Struct. Funct. Genet. 28:117–130.

19. Hjorth, A., F. Carriere, C. Cudrey, H. Woldike, E. Boel, D. M. Lawson, F.

Ferrato, C. Cambillau, G. G. Dodson, L. Thim, and R. Verger. A structural

domain (the lid) found in pancreatic lipases is absent in the guinea pig (phospho)

lipase. Biochemistry. 32:4702-4707.

20. Hutcheon, G. W., N. Vasisht, and A. Bolhuis. 2005. Characterisation of a highly

stable alpha-amylase from the halophilic archaeon Haloarcula hispanica.

Extremophiles. 9:487-495

21. Jaeger, K-E., B. W. Dijkstra, and M. T. Reetz. 1999. Bacterial biocatalysts:

molecular biology, three-dimensional structures, and biotechnological applications

of lipases. Annu. Rev. Microbiol. 53:315–351.

22. Jaeger, K. E. and M. T. Reetz. 1998. Microbial lipases form versatile tools for

biotechnology. Trends Biotechnol. 16:396-403.

23. Jaenicke, R., and G. Bohm. 1998. The stability of proteins in extreme

environments. Curr. Opin. Struct. Biol. 8:738-48.

101

24. Jeong, S. T., H. K. Kim, S. J. Kim, S. W. Chi, J. G. Pan, T. K. and S. Eryu. Novel

zinc-binding center and a temperature switch in the Bacillus stearothermophilus L1

lipase. J. Biol. Chem. 277:17041-7.

25. Khyami-Horani, H. 1996. Thermotolerant strain of Bacillus licheniformis producing

lipase. World J. Microbiol. Biotechnol. 12: 399-401.

26. Kim, M. H., H. K. Kim, J. K. Lee, S. Y. Park, and T. K. Oh. 2000. Thermostable

lipase of Bacillus Stearothermophilus: high-level production, purification, and

calcium-dependent thermostability. Biosci. Biotechnol. Biochem. 64:280-6.

27. Kim, H. K., S. Y. Park, J. K. Lee, and T. K. OH. 1998. Gene cloning and

characterization of thermostable lipase from Bacillus stearothermophilus L1. Biosci.

Biotechnol. Biochem. 62: 66-71.

28. Lang, D., B. Hofmann. L. Haalck. H. J. Hecht. F. spener, R. D. Schmid, and D.

Schomburg. 1996. Crystal structure of a bacterial lipase from Chromobacterium

viscosum ATCC 6918 refined at 1.6 angstroms resolution. J. Mol. Biol. 259:704-

717.

29. Lee, P., G. H. Chung, and J. S. Rhee. 1993. Purification and characterization of

Pseudomonas fluorescens SIK-W1 lipase expressed in Escherichia coli. Biochim.

Biophys. Acta. 1169:156–164.

30. Lee, W., Y. S. Koh, K. J. Kim, B. C. Kim, H. J. Choi, D. S. Kim, M. T. Suhartono

and Y.R. Pyun. 1999. Isolation and characterization of a thermophilic lipase from

Bacillus thermoleovorans ID-1. FEMS Microbiol. Lett. 179:393–400.

102

31. Lesuisse, E., K. Schanck, and C. Colson. 1993. Purification and preliminary

characterization of the extracellular lipase of Bacillus subtilis 168, an extremely

basic pH-tolerant enzyme. Eur. J. Biochem. 216:155-60.

32. Macedo-Ribeiro, S., B. Darimont, R. Sterner, and R. Huber. 1996. Small

structural changes account for the high thermostability of 1[4Fe–4S] ferredoxin

from the hyperthermophilic bacterium Thermotoga maritima. Structure. 4:1291–

1301.

33. Martinez, C., P. De Geus, M. Lauwereys, G. Matthyssens, and C. Cambillau.

1992. Fusarium solani cutinase is a lipolytic enzyme with a catalytic serine

accessible to solvent. Nature. 356:615-618.

34. Matsumae, H., and T. Shibatani. 1994. Purification and characterization of lipase

from Serratia marcescens Sr41 8000 responsible for asymmetric hydrolysis of 3-

phenylglysidic acid esters. J. Ferment. Bioeng. 77: 152–158.

35. Nardini, M., D. A. Lang, K.E. Jaeger and B.W. Dijkstra. 2000. Crystal structure

of Pseudomonas aeruginosa lipase in the open conformation. The prototype for

family I.1 of bacterial lipases. J. Biol. Chem. 275: 31219–31225.

36. Nawani, N., and J. Kaur. 2000. Purification, characterization and thermostability of

lipase from a thermophilic Bacillus sp. J33. Mol. Cell Biochem. 206:91-96.

37. Noble, M. E., A. Cleasby, L. N. Johnson, M. R. Egmond, and L. G. Frenken.

1993. The crystal structure of triacylglycerol lipase from Pseudomonas glumae

reveals a partially redundant catalytic aspartate. FEBS Lett. 331:123-128.

103

38. Rashid, N., Y. Shimada, S. Ezaki, H. Atomi, and T. Imanaka. 2001. Low-

temperature lipase from psychrotrophic Pseudomonas sp. strain KB700A. Appl.

Environ. Microbiol. 67:4064-9.

39. Rathi, P., S. Bradoo, R. K. Saxena, and R. Gupta. 2000. A hyper-thermostable,

alkaline lipase from Pseudomonas sp. With the property of thermal activation.

Biotech. Lett. 22: 495-498.

40. Rosenstein, R., and F. Gotz. 2000. Staphylococcal lipases: biochemical and

molecular characterization. Biochimie. 82:1005-14.

41. Rua Luisa, M., C. Schmidt-Dannert, S. Wahl, A. Sprauer, and R. D. Schmid.

1997. Thermoalkalophilic lipase of Bacillus thermocatenulatus large-scale

production, purification and properties: aggregation behaviour and its effect on

activity. J. Biotech. 56: 89-102.

42. Russell, R. J., J. M. Ferguson, D. W. Hough, M. J. Danson, and G. L. Taylor.

1997. The crystal structure of citrate synthase from the hyperthermophilic

archaeon Pyrococcus furiosus at 1.9 resolution. Biochemistry. 36:9983-9994.

43. Sambrook. J., E. F. Fritsch and T. Maniatis. 1989. Molecular cloning: A

laboratory manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New

York.

44. Sarda, L., and P. Desnuelle. 1958. Action of pancreatic lipase on emulsified

esters. Biochim. Biophys. Acta. 30:513-521.

45. Schmidt-Dannert, C. 1999. Recombinant microbial lipases for biotechnological

applications. Bioorg. Med. Chem. 7:2123-2130.

104

46. Schmidt-Dannert, C., H. Sztajer, W. Stocklein, U. Menge and R. D. Schmid.

1994. Screening, purification and properties of a thermophilic lipase from Bacillus

thermocatenulatus. Biochim. Biophys. Acta. 1214: 43–53.

47. Schmidt-Dannert, C., M. L. Rua, H. Atomi, and R. D. Schmid. 1996.

Thermoalkalophilic lipase of Bacillus thermocatenulatus. I. molecular cloning,

nucleotide sequence, purification and some properties. Biochim. Biophys. Acta.

1996 1301:105-114.

48. Simons, J. W., M. D. van Kampen, I. Ubarretxena-Belandia, R. C. Cox, C. M.

Alves dos Santos, M. R. Egmond, and H. M. Verheij. 1999. Identification of a

calcium binding site in Staphylococcus hyicus lipase: generation of calcium-

independent variants. Biochemistry. 38:2-10.

49. Sinchaikul, S., B. Sookkheo, S. Phutrakul, F. M. Pan, and S. T. Chen. 2001.

Optimization of a thermostable lipase from Bacillus stearothermophilus P1:

overexpression, purification, and characterization. Protein Expr. Purif. 22:388-398.

50. Snellman, E. A., E. R. Sullivan, and R. R. Colwell. 2002. Purification and

properties of the extracellular lipase, LipA, of Acinetobacter sp. RAG-1. Eur. J.

Biochem. 269:5771-5779.

51. Spencer, D. S., K. Xu, M. T. Logan and H. X. Zhou. 2005. Effects of pH, salt, and

macromolecular crowding on the stability of FK506-binding protein: an integrated

experimental and theoretical study. J. Mol. Biol. 351:219-232.

52. Svetlitshnyi, V., F. Rainey, and J. Wiegel. 1996. Thermosyntropha lipolytica gen.

nov., sp. nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing

105

106

short and long chain fatty acids in syntrophic coculture with a methanogenic

archaeum. Int. J. Syst. Bacteriol. 46:1131-1137.

53. Tanner, J. J., R. m. Hecht, and K. L. Krause. 1996. Determinants of enzyme

thermostability observed in the molecular structure of Thermus aquaticus D-

glyceraldehyde-3-phosphate dehydrogenase at 2.5 Å resolution. Biochemistry.

35:2597–2609.

54. Tyndall, J. D., S. Sinchaikul, L. A. Fothergill-Gilmore, P. Taylor, and M. D.

Walkinshaw. 2002. Crystal structure of a thermostable lipase from Bacillus

stearothermophilus P1. J. Mol. Biol. 323:859-69.

55. Verger, R. and G. H. De Haas. 1976. Interfacial Enzyme Kinetics of Lipolysis.

Annu. Rev. Biophys. Bioeng. 5:77-117.

56. Villeneuve, P., J. M. Muderhwa, J. Graille, and M. J. Haas. 2000. Customizing

lipases for biocatalysis: a survey of chemical, physical and molecular biological

approaches. J. Mol. Catal. B Enzym. 9:113–148.

57. Wiegel, J. 1998. Anaerobic alkalithermophiles, a novel group of extremophiles.

Extremophiles. 2:257-267.

58. Winkler, U. K., and M. Stuckmann. 1979. Glycogen, hyaluronate, and some other

polysaccharides greatly enhance the formation of exolipase by Serratia

marcescens. J. Bacteriol. 138:663-70.

107

Step Total volume

(ml)

Total activity

(units)

Total protein

(mg)

S. activity

U· mg-1

Yield % Purification

factor

Supernatant 22000 2900 28600 0.1 100 1

Holofiber 10KDa cutoff 5000 2640 3250 0.81 91 8

(NH4)2SO4 precipitation 800 1580 392 4.0 54.5 40

Octyl sepharose LipA 300 660 47 14 23 140

Octyl sepharose LipB 210 390 29 13.4 13.4 134

Table 2.1. Purification of LipA and LipB produced extracellularly by Thrmosyntropha lipolytica.

Table 2.2. Effect of various metal ions and inhibitors on lipase activity. Relative remaining activity (%) LipA LipB 1mM 10mM 1mM 10mM

Caa 100 ±5 100 ±5 100 ±5 100 ±5 Mg 95 ±5 65 ±5 97 ±5 73 ±5 Fea 12 ±5 0 20 ±5 0 Mna, b 340 ±5 270 ±15 315 ±15 250 ±15 K 65 ±5 30 ±10 85 ±5 70 ±10 Zn 50 ±15 44 ±15 62 ±15 36 ±15 Na 78 ±5 70 ± 5 83 ±5 80 ±5 Cs 77 ±5 37 ±10 72 ±5 44 ±5 Cu 45 ±10 0 56 ±10 0 Al 28 ±5 0 30 ±5 0 Ni 80 ±5 22 ±5 50 ±5 15 ±5 Co 60 ±5 60 ±5 68 ±5 49 ±5 PMSF 70 ±15 6 ±5 62 ±15 0 EDTA 60 ±15 11 ±5 81 ±5 0

a Assays were conducted with triolein and lipase activity quantified by NEFA C kit. b The addition of Mn or Mn and Ca together gave the same value

108

FIG. 2.1.

Extracellular lipase activity during growth at pH60°C 7.6 (A) and pH60°C 9.0 (B), O.D.

(■), specific activity (■).

109

110

FIG. 2.2.

Purified LipA and LipB proteins were analyzed by SDS–PAGE. Lane 1, Molecular

weight Marker (kDa); lane 2, Purified LipB after Octyl Sepharose; lane 3, purified LipA

after Q-Sepharose Fast Flow chromatography. Gels were stained with GelCode® Blue

Stain Reagent (PIERCE).

111

112

FIG. 2.3.

Effect of temperature on LipA (○) and LipB (■) activity. The lipase assay was

conducted as mentioned in the materials and methods section. The substrate used for

the assay was pNPL. Similar temperature profile but lower specific activity was also

obtained when pNPL was substituted with pNPP. The inside window is the temperature

profile when only TAPS buffer was used at pH 9.0. A 100% relative activity was 11.8

±0.5 U·mg-1 and 13.0 ±0.6 U·mg-1 for LipA and LipB, respectively. Activity values were

corrected from controls without enzyme.

113

114

FIG. 2.4.

The effect of pH on activity (A), LipA (○) and LipB (■), and stability (B), LipA (open

symbols) and LipB (closed symbols). Mixed buffer of 100mM of HEPES, TAPS and

CAPS was used and adjusted at room temperature. 100% relative activity was 12.4

±0.5 U·mg-1 and 12.8 U·mg-1 ±0.3 U·mg-1 for LipA and LipB, respectively. For stability

experiments (C), 100 mM of the following buffers were used: glycine–HCl (▲), sodium

acetate buffer (♦) MES buffer (■),TAPS buffer (●), CAPS buffer (▀). Results are

expressed as percentage of maximal activity. The lipase assays were conducted as

described in the materials and methods section.

115

116

FIG. 2.5.

Thermostability of LipA (A) and LipB (B). The thermostability of the lipases in pH 8.0

buffer was tested by measuring the residual activity after incubation at various

temperatures; 100 ºC (■), 75 ºC (○) and 60 ºC (∆). All measurements were in triplicate.

The lipase assay was conducted as discussed in the methods section. A 100% relative

activity is equal to the specific activity of the enzyme before incubation, which is 13.0

±0.6 U·mg-1 and 13.6 U·mg-1 ±0.6 for LipA and LipB, respectively.

117

118

FIG. 2.6.

The activity of purified LipA (□) and LipB (■) toward substrates with different acyl

chain lengths was determined under standard conditions using p-nitrophenyl

esters (A) and triglycerides (B). A 100% relative activity is 12.4 ±0.5 U·mg-1 and 13.0

±0.6 U·mg-1 for LipA and LipB, respectively.

119

120

FIG. 2.7.

Positional specificity of purified LipA and LipB. The specificity was resolved on

silica-TLC plate after 8 hours assay. Lanes:1, triolein; 2, 1,3-diolein; 3, 1,2 diolein; 4, 1-

monoolein; 5, 2-monoolein; 6, hydrolysis products of LipA purified after octyl sepharose;

7, hydrolysis products of LipA purified after Q sepharose; 8, hydrolysis products of pure

LipB; 9, control (no enzyme was added).

121

122

CHAPTER 3

EFFECTS OF VARIOUS DETERGENTS ON TWO ALKALITHERMOPHILIC LIPASES

FROM THERMOSYNTROPHA LIPOLYTICA1

1Salameh, M. and J. Wiegel. 2006. To be submitted to FEBS Journal.

123

Abstract

Thermosyntropha lipolytica DSM 11003, an anaerobic thermophilic lipolytic bacterium,

produces two highly thermoalkaliphilic and thermostable lipases, LipA and LipB.

Nonionic, anionic and cationic detergents affected the activity and the stability of both

lipases. All nonionic detergents exhibited activation when used below their critical

micelle concentration (CMC) values, but caused a slight inhibition above their CMC.

Anionic detergents showed activation only at high concentration exceeding their CMC

values. Cationic detergent CTAB was inhibitory for both lipases. In the absence of

detergents, the vmax of both LipA and LipB were 12.4 U·mg-1 and 13.3 U·mg-1 and K0.5

were 1.8 mM and 1.65 mM, respectively at 96°C. In the presence of 0.2% SDS in the

assay the vmax values increased to 105 U·mg-1 and 112 U·mg-1, and K0.5 values

decreased to 800 µM and 740 µM for LipA and LipB, respectively. Assay analyses using

diisopropyl p-nitrophenylphosphate (E600) with increasing concentration of SDS and

Tween 20 strongly suggest that SDS and Tween 20 do bind to the lid domain and/or

active site pocket, thus promoting conformational changes which lead to the

displacement of the lid that covers the active site, so that the site becomes accessible to

the substrate.

124

Introduction

Lipases (E.C. 3.1.1.3) are carboxyl ester hydrolases that catalyze the hydrolysis

and synthesis of long chain acylglycerols [1, 2]. Despite the remarkable tertiary

structural homology of the enzymes obtained from distantly related organisms, there is

little sequence homology among lipases in general [3]. Furthermore, the specific

characteristics of lipases such as size, amino acid sequences, substrate specificity,

thermal activity, stability under various conditions and enantioselectivity vary widely,

depending primarily on the source of the enzyme. Lipases are activated at the water-

lipid interface; they show little activity when the substrate is in the monomeric form and

activity increases dramatically above the solubility limit, where they start to form

emulsions [1, 2]. This fact has led to the emergence of a phenomenon known as

interfacial activation [2, 4].

Triton X-100, Tween 20 and Tween 80 are nonionic polyoxyethylene detergents.

The hydrophobic part usually consists of an alkyl chain (branched or unbranched), and

the hydrophilic part is made up of uncharged ethylene oxide units. Aqueous solutions of

the nonionic polyoxyethylene detergents form two liquid phases upon temperature

increase. One of these phases is detergent-enriched and called the coacervate phase,

whereas the other is detergent-depleted [5]. Terstappen et al. [6] found that there is a

positive correlation between protein hydrophobicity and its partitioning into the

coacervate phase which confirmed that protein-detergent interactions in such systems

are primarily hydrophobic.

One thousand tons of lipases are needed every year for detergent industry [4].

For a lipase to be useful as a detergent additive, it should be alkaliphilic, thermostable,

125

resistant to denaturation in the harsh detergent environment, and active in the presence

of detergent. High levels of thermal stability are positively correlated with stability in the

presence of other denaturing agents, including detergents and organic solvents, which

suggest that molecular mobility/flexibility is the prime determinant of susceptibility to

irreversible denaturation [11]. Enzymes from thermophilic organisms are generally more

robust, more thermo and organo stable, and active at elevated temperatures [12-15].

T. lipolytica is an anaerobic thermophilic organotrophic lipolytic alkalitolerant bacterium.

It can grow on triglycerides and ferment long chain fatty acids only in syntrophic co-

culture with Methanobacterium strain JW/VS-M29, but does not utilize the liberated

glycerol [16]. Previously, we reported the purification and characterization of two highly

thermophilic lipases, LipA and LipB [17]. They are the most thermophilic and

thermostable lipases reported so far: maximum catalytic activity observed at 96°C, and

optimum pH80°C around 8.5, and retaining 50% activity after 20 hours incubation at 75°C.

The optimum activity is observed using long chain fatty acids glycerides as substrates

[17]. Here, we describe and discuss the effects of different detergents on the enzymatic

activity, thermal stability and structural implications.

Results and Discussion

SDS effect on temperature optima. Activity of the electrophoretically-pure lipases LipA

and LipB were analyzed in the absence of SDS using a 20 min assay. LipA has a broad

maximum activity range between 86 °C to 96 °C, whereas LipB has its highest activity

around 96 °C [17]. However, the addition of 0.2% SDS (final concentration) increased

the specific activity of LipA from 12.2 U·mg-1 to 105 U·mg-1, with maximum catalytic

activity observed in the range of 90 °C and 96 °C. LipB specific activity was increased

126

from 13.0 U·mg-1 to 112 U·mg-1, and maximum activity was shifted down from 96 °C to

92 °C (Fig. 3.1). Similar shifts were observed when TAPS was used as the assay buffer

[17]. The detergents Tween 20 and Triton X-100 did not produce any noticeable shift in

temperature profiles. A similar effect of SDS on LipA and LipB catalytic activity was

observed for the lipase TIL from the temperature tolerant Thermomyces lanuginosus [7].

All of these enzymes are alkalithermophilic, more thermostable than most lipases and

are activated by SDS. Another enzyme from sun flower, phospholipase D, was found to

be activated by 10 times in the presence of SDS [18].

Substrate hydrolysis rates. The hydrolysis rates of p-nitrophenyl laurate in the

absence and presence of SDS were elucidated (Fig. 3.2). All substrate saturation

curves were sigmaoidal, i. e., when the substrate exceeded its solubility limit, the

hydrolysis rate increased non hyperbolically. These results suggest that LipA and LipB

are true lipases that show little activity toward soluble substrate and are interfacially

activated by pNPL. The substrate hydrolysis profile of sunflower phospholipase D

showed interfacial activation and S shaped hydrolysis profile, it was converted into a

hyperbolic like curve in the presence of SDS and Triton X-100 (18).

The vmax of both LipA and LipB calculated from the hydrolysis rate curves were

12.6 U·mg-1 and 13.3 U·mg-1, K0.5 were 1.8 mM and 1.65 mM, respectively. In the

presence of 0.2% SDS, LipA and LipB behaved more like esterases, where the lag

phase that was observed initially without SDS was greatly reduced. The vmax values

increased to 105 U·mg-1 and 112 U·mg-1 and K0.5 values decreased to 800 µM and 740

µM for LipA and LipB, respectively. The hydrolysis rates of both enzymes were

127

proportional to the amount of the enzyme used at both below and above the solubility

limit of pNPL.

Effect of detergents on activity. Detergents in general have close resemblance to

lipase substrates, they form aggregates in aqueous solution when exceeding certain

concentration. Therefore, it is expected that detergents will influence lipase activity. We

investigated the impact of different detergents on enzymatic activities of LipA and LipB

by recording specific activity profiles at different detergent concentrations using pNPL

as substrate (Fig. 3.3). There have been a few studies involving the effect of detergents

on lipase activity [7, 18, 19], making it possible to compare our data with previously

published results. However, all these studies used p-nitrophenyl butyrate as substrate.

This is the first study that involves thermophilic lipases and p-nitrophenyl laurate as

substrate.

Among all tested detergents, SDS was found to have the greatest impact on

catalytic activity of both lipases (Fig. 3.3A). The effect of SDS on activity started at

premicellar concentration of 1 mM, but the activity was only 3-6 U·mg-1 higher than

when tested in buffer alone. The activity was drastically increased at the CMC point

around 2 mM (Table 3.1) and peaked at 7 mM (0.2%) SDS, where the activity was

recorded at 104 and 109 U·mg-1 for LipA and LipB, respectively. At higher SDS

concentrations, the activity declined to reach 4.5 U·mg-1 and 10.6 U·mg-1 for LipA and

LipB, respectively. This result suggested that LipA was (partially) unfolded by SDS,

whereas LipB is more robust and resistant to high SDS concentrations.

A recent study by Mogensen et al. [7] showed a detailed characterization of the

effect of various detergents on the enzymatic activity and thermal stability of

128

Thermomyces lanuginosus lipase (TlL). LipA and LipB are reminiscent to TIL in being

thermoalkaliphilic lipases, thermostable and activated by SDS.

These results are in agreement with those of Mogensen et al. [7], where SDS

was found to activate T. lanuginosus lipase (TlL) around the CMC level. Above the CMC

level, the activity declined but still significantly higher than without detergent. Although

TIL activity was greatly reduced at higher SDS concentrations (reduced from 100 at

CMC level to 10 folds activity), but was never inhibited as observed with LipA and LipB.

Substrate and detergent interactions with lipases can be complex and

unpredictable [7-10]. However, there is behavior similarity between T. lipolytica lipases

and TIL toward various detergents and especially SDS. The degree of activation and

inhibition could be quite different; however both are well described among all of them.

In case of the other ionic detergent sodium cholate, we didn’t observe the same

activation/inhibition magnitude effect as of SDS. At monomeric and miceller

concentrations the activity increase did not exceed more than 4U·mg-1, indicating that

SDS binding is more effective (Fig. 3.3B).The cationic detergent CTAB was inhibitory to

LipA and to a less extent LipB (Fig. 3.3C). This might be due to unfavorable electrostatic

interactions that might cause unfolding of the enzyme [20] and/or disrupt substrate

binding.

All the non ionic detergents, Triton X-100, Tween 20, and Tween 80, behaved

similarly in their effect on LipA and LipB catalytic activity (Fig 3.3D, E, F). Activation

occurs at concentration below their CMC levels (Table 2.1) followed by a gradual

decline in activity to a plateau that is similar to the one without added detergent, an

inhibition was only observed with Triton X-100. The similarity of activity profiles between

129

non ionic detergents and ionic ones based on the fact that the activation started at low

concentrations below their CMC values and therefore does not involve micelles. Similar

results were also observed by others [7, 9]. However, in case of SDS the activation

started in the presence of detergent monomers, but maximal activation as well as

inhibition occurred in the presence of micelles. The possible explanation for this result is

that SDS micelles bind preferably to the lid and activate the enzyme by triggering

conformational changes, and that detergent binding and substrate binding cannot be

identical. This conclusion is supported by other studies using fluorescence and X-ray

crystallography which showed that micelles indeed bind to the lid that covers the active

site and promote conformational changes [7, 10, 21, 22]. The continuous decrease in

enzyme activity with increasing detergent concentration could be attributed to unfolding

of the enzymes by SDS.

SDS and Tween 20 promote conformational changes. To examine whether the

detergent micelles do bind to the lid region of LipA and LipB and trigger conformational

changes, diisopropyl p-nitrophenylphosphate (E600) was used. E600 is a serine

inhibitor that can covalently bind to the serine in the active site and irreversibly inhibit

the lipase [23]. It is believed that E600 has access to the active site only when the

enzyme is in the open conformation. Therefore, if the micelles bind to the lid and trigger

conformational changes that expose the active site, E600 will have easy access to the

serine and inhibits the enzyme [7, 10]. The results in table 3.2 show that E600 did

indeed inhibit both lipases in the presence of SDS micelles. This experiment

demonstrated micelles promote conformational changes by binding to the lid/active site

domain mimicking the action of actual substrate. As observed, the activation of LipA and

130

LipB started at low concentration of SDS below the CMC value. In addition, at 1 mM

SDS we observed inhibition of both enzymes in the presence of E600, which means

detergent molecules, may bind as monomers on protein’s hot spots [7, 24]. The

hydrophobic moieties such as alkyl chains in the non ionic detergents are likely to bind

to the active site of the lipase because of the high physical similarity to lipase substrates

[7, 9].

To find out if non ionic detergents bind to the enzyme, we incubated the enzymes

with E600 along with increasing concentrations of Tween 20 (Table 2.2). If Tween 20

binds to the enzyme, it will trigger conformational changes that would convert the

enzyme from closed to open conformation allowing accessibility of E600 to bind to the

active site. The results in Table 3.2 demonstrate that the non ionic detergent does

promote conformational changes as the inhibition increased as a function of detergent

concentration. Hermoso et al. showed by means of crystallography that a non ionic

detergent activated the porcine lipase and bound tightly to the active site pocket, acting

like a substrate analog. However, at submicellar detergent concentration, the enzyme

activity was inhibited [10]. The interactions between detergents and lipases are primarily

hydrophobic. However, the charged groups of cationic and anionic detergents play an

important contribution to this interaction, therefore a considerable divergence on the

behavior of cationic, anionic and as well as nonionic detergents with lipases could be

seen [9]. The question here is if E600 inhibits LipA and LipB, why is complete inhibition

not observed? LipA and LipB are highly thermostable and thermophilic enzymes, and

cold labile [17], which suggest these enzymes have very rigid confirmation that play

important role in the activation, inhibition processes. This fact is also supported by the

131

inability of E600 to inhibit both enzymes when enzymes, detergent and 600 were

incubated at room temperature instead of 60 °C (Table 3.2).

Detergent effect on stability. To examine the stability of both enzymes in the presence

of non ionic and an ionic detergents. Aliquots of the purified enzymes were incubated at

room temperature with 1% detergent. At room temperature, Tween 20 has no effect on

the stability of either enzymes as compared to the control; both enzymes retained 95%

activity after 48 hours incubation (Fig. 3.4). In the presence of SDS however, the

stability was gradually reduced, LipA was the most affected by SDS as the activity was

reduced to 6 U·mg-1 while LipB activity was reduced to 8 U·mg-1 after 48 hours

incubation. The nonionic detergents are considered mild detergents; they do not interact

extensively with the protein surface, whereas ionic detergents such as SDS, are more

aggressive and highly reactive resulting in slowly unfolding the protein [25]. The slow

rate of unfolding in the presence of SDS is probably attributed to the rigid structure and

the high stability of the enzymes.

Materials and Methods

Culture and growth conditions. Thermosyntropha lipolytica was grown in a basal

medium containing 0.75% yeast extract as carbon and energy sources under nitrogen

gas phase. The basal medium contained (per liter) 0.3 g of K2HPO4, 0.3 g of KCl, 0.5 g

of NaCl, 1.0 g of NH4Cl, 0.1 g of MgCl2·6H2O, 0.02 g of CaCl2·2H2O, 3.0 g of NaHCO3,

3.0 g of Na2CO3, 0.5 g of Na2S·9H2O, 0.15 g of cysteine, 2 ml of vitamin solution and

2.5 ml of trace element solution [26]. The pH of the medium was adjusted at 8.225°C and

the growth temperature was at 60 °C.

132

Lipase assay. Lipase assays were carried out as using a spectrophotometric method

with the chromogenic substrates p-nitrophenyl laurate (pNPL) and p-nitrophenyl

palmitate (pNPP) (Sigma) [27]. Reaction mixture contained 100 µl of enzyme solution,

25 µM MnCl2, 1,088 µl of freshly prepared buffer containing 100 mM HEPES, 100 mM

TAPS and 100 mM CAPS buffer (Sigma), and 12 µl of 300 mM pNP-laurate or pNP-

palmitate in acetonitrile (final concentration in the assay, 3 mM). The assay was carried

out for 20 minutes at 96 °C. The reaction mixture was then cleared by centrifugation and

the liberated p-nitrophenol was determined at A405. One unit is defined as the amount of

the enzyme catalyzing the release of 1 µmol of p-nitrophenol (ε=1.82 × 104 M-1cm-1) per

min from pNP-laurate/palmitate.

Employing these conditions, the kinetics of pNPL hydrolysis was linear as a function of

time (over 40 minutes) and enzyme concentration (5-50 µg). The value of every assay

was corrected of substrate hydrolysis of an enzyme free blank under the same test

conditions.

Purification of lipases. Purification of LipA and LipB was conducted as described

previously [17]. In summary, the two lipases were purified from 20 liter culture

supernatant through filtration by using 10 kDa Amicon hollow fiber (Millipore), purified

from supernatant by stepwise ammonium sulfate precipitation and hydrophobic

interaction chromatography, LipA and LipB eluted at 1.8 M and 0.5 M (NH4)2SO4,

respectively, and then desalted and concentrated by using filter membranes (Millipore).

Gel electrophoreses. Electrophoretic analyses were performed with a Bio-Rad Mini-

Protean II cell unit, at room temperature. SDS/PAGE and non denaturing PAGE were

133

performed essentially as described by Sambrook and Ausubel [28, 29]. Gels were

stained with GelCode® Blue Stain Reagent (Pierce).

Protein concentration. Protein concentrations were routinely determined with the

bicinchoninic acid (BCA) protein assay kit (Pierce) following the manufacturer’s

instructions.

Determination of critical micelle concentration (CMC). The CMC values of

detergents were determined using the fluorescence properties of N-phenyl-1-

naphthylamine (NPN) as described elsewhere [7, 30, 31]. Basically, in aqueous solution

NPN has a very low fluorescence quantum; however, the partitioning of NPN into an

apolar environment such as micelles is associated with a dramatic increase in the

fluorescence intensity of the probe. Detergents were suspended in the assay buffer at

room temperature at varying concentrations (1 µM to 25 mM) and then incubated for 1 h

in the presence of 10 µM N-phenyl-1-naphthylamine (NPN) at 50 °C. The fluorescence

intensity was measured using a computerized fluorometer (Shimadzu RF-5301PC) (λex

= 350 nm, λem = 435 nm, 1 nm bandwidth). When plotting NPN fluorescence as a

function of detergent concentration, the CMC was determined from the breakpoint of the

curve.

The effect of SDS on enzymatic activity. The temperature and pH optima for

maximum catalytic activity of LipA and LipB were previously determined along with a

detailed thermostability study [17]. A temperature gradient incubator was used to

determine the temperature activity profiles of both enzymes in the presence of 0.2%

SDS. The enzyme assay was performed at pH25°C 9.4 and 9.6 which are equal to

pH80°C 8.5 and 8.8 for LipA and LipB, respectively. The enzyme assays were performed

134

as described above, using as a buffer 100 mM HEPES, 100 mM TAPS and 100 mM

CAPS. After the addition of substrate and SDS, the assay solution was shaked for 5

minutes at 85 °C, vortexed for 10 seconds and 10 µg of purified enzyme was added.

The assay duration was 15 minutes after which the reactions were promptly stopped in

ice. Assays were conducted in triplicate.

The effect of various detergents on activity and stability. All detergents were from

Sigma, USA. Sodium dodecyl sulfate (SDS) and sodium cholate are anionic detergents,

hexadecyltrimethylammonium bromide (CTAB) is cationic detergent,

t-Octylphenoxypolyethoxyethanol (Triton X-100), polyoxyethylene sorbitan monolaurate

(Tween 20) and polyoxyethylene sorbitan monooleate (Tween 80) are non ionic

detergents. The standard assay as described earlier was employed with increasing

concentration of detergent. The assay temperature was 92 °C, which was a

compromise between lipase activity and substrate stability in the presence of detergent.

To determine detergent effect on enzyme stability at room temperature, aliquots of

purified LipA and LipB were incubated in the presence of 1% detergent in sealed 2 ml

serum bottles, which were specified for every time point. Each sample contained 15µg

of protein in 0.6 ml of 100 mM Tris buffer, pH 8.0 and 0.5 M ammonium sulfate. After

incubation, the samples were concentrated and desalted by using 10 kDa cut off

centricon filter tubes (Millipore), protein quantified as mentioned earlier and assayed for

lipase activity. The assays were in triplicate and product formation was proportional to

the incubation time above 40 min.

Inhibition of LipA and LipB by diisopropyl p-nitrophenylphosphate (E600).

Enzymatic activity experiments in the presence of the inhibitor E600 were performed as

135

described previously [7, 10] with the following modification. 10 µg of purified LipA and

LipB were incubated at 50 °C in 100 mM TAPS buffer pH 9.0 in the presence of 5 mM

E600 and increasing concentrations of SDS and Tween 80 for 120 minutes, final

volume was 300 µl. E600 was excluded from the controls. At the end of the incubation,

enzyme assays were performed as described earlier. Each assay was carried out in

triplicate.

Acknowledgment

The authors are thankful to Dr. Timothy Davis for his help in fermentation and

conducting flourometer measurements.

136

References

1. Sarda, L & Desnuelle P (1958) Action of pancreatic lipase on emulsified esters.

Biochim Biophys Acta 30, 513-521.

2. Ferrato F, Carriere F, Sarda L & Verger R (1997) A critical reevaluation of the

phenomenon of interfacial activation. Methods Enzymol 286, 327-347.

3. Bell PJL, Nevalainen H, Morgan HW & Bergquist PL (1999) Rapid cloning of

thermoalkalophilic lipases from Bacillus sp. Using PCR. Biotech. Lett 21, 1003-1006.

4. Jaeger KE & Reetz MT (1998) Microbial lipases form versatile tools for

biotechnology. Trends Biotechnol 16, 396-403.

5. Nakagawa & Nakagawa (1966) Solubilization In: M.J. Schick, Editor, Nonionic

Surfactants, Marcel Dekker, New York, 558–603.

6. Terstappen GC, Ramelmeier RA & Kula MR (1993) Protein partitioning in detergent-

based aqueous two-phase systems. J Biotechnol 28, 263–275.

7. Mogensen JE, Sehgal P & Otzen DE (2005) Activation, Inhibition, and destabilization

of Thermomyces lanuginosus lipase by detergents. Biochemistry 44, 1719 -1730.

8. Misiorowski RL & Wells MA (1974) The activity of phospholipase A2 in reversed

micelles of phosphatidylcholine in diethyl ether: Effect of water and cat ions.

Biochemistry. 13, 4921–4927.

9. Helistö P & Korpela T (1998) Effect of detergents on activity of microbial lipases as

measured by the nitrophenyl alkanoate esters method. Enzyme Microb Technol 23,

113-117.

10. Hermoso J, Pignol D, Kerfelec B, Crenon I, Chapus C, & Fontecilla-Camps J. C.

(1996) Lipase activation by nonionic detergents. The crystal structure of the porcine

137

lipase-colipase-tetraethylene glycol monooctyl ether complex, J Biol Chem 271,

18007-18016.

11. Cowan DA (1997) Thermophilic proteins: stability and function in aqueous and

organic solvents. Comp Biochem Physiol A Physiol 118,429-438.

12. Ejima K, Liu J, Oshima Y, Hirooka K, Shimanuki S, Yokota Y, Hemmi H, Nakayama

T & Nishino T (2004) Molecular cloning and characterization of a thermostable

carboxylesterase from an archaeon, Sulfolobus shibatae DSM5389: Non-linear

kinetic behavior of a hormone-sensitive lipase family enzyme. J Biosci Bioeng 98,

445-451.

13. Pantazaki AA, Pritsa AA & Kyriadidis DA (2002) Biotechnologically relevant

enzymes from Thermus thermophilus. Appl Microbiol Biotechnol 58, 1-12.

14. Fucinos P, Dominguez A, Sanroman MA, Longo MA, Rua ML & Pastrana L (2005)

Production of thermostable lipolytic activity by Thermus species. Biotechnol Prog 21,

1198-1205.

15. Li H & Zhang X (2005) Characterization of thermostable lipase from thermophilic

Geobacillus sp. TW1. Protein Expr Purif 42,153-159.

16. Svetlitshnyi V, Rainey F & Wiegel J (1996) Thermosyntropha lipolytica gen. nov., sp.

nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing short and

long chain fatty acids in syntrophic coculture with a methanogenic archaeum. Int J

Syst Bacteriol 46, 1131-1137.

17. Salameh M & Wiegel J (2006) Purification and characterization of two highly

thermophilic alkaline lipases from Thermosyntropha lipolytica. Appl Environ Microbiol

(submitted).

138

18. Abousalham A, Nari J, Teissere M, Ferte N, Noat G & Verger R (1997) Study of fatty

acid specificity of sunflower phospholipase D using detergent/phospholipid micelles

Eur J Biochem 248, 374-379.

19. Lai DT & O'Connor CJ (2000) Synergistic effects of surfactants on kid pregastric

lipase catalyzed hydrolysis reactions. Langmuir 16, 115 -121.

20. Otzen DE (2002) Protein unfolding in detergents: Effect of micelle structure, ionic

strength, pH, and temperature. Biophys J 83, 2219-2230.

21. Yapoudjian S, Ivanova MG, Brzozowski AM, Patkar SA, Vind J, Svendsen A &

Verger R (2002) Binding of Thermomyces (Humicola) lanuginosa lipase to the mixed

micelles of cis-parinaric acid/NaTDC. Eur J Biochem 269, 1613-1621.

22. Cajal Y, Svendsen A, Girona V, Patkar SA & Alsina MA (2000) Interfacial control of

lid opening in Thermomyces lanuginosa lipase. Biochemistry 39, 413-423.

23. Maylie MF, Charles M & Desnuelle P (1972) Action of organophosphates and

sulfonyl halides on porcine pancreatic lipase. Biochim Biophys Acta 276, 162-175.

24. Ibel K, May RP, Kirschner K, Szadkowski H, Mascher E & Lundahl P (1990) Protein-

decorated micelle structure of sodium-dodecyl-sulfate-protein complexes as

determined by neutron scattering. Eur J Biochem 190, 311-318.

25. Stobiecka A, Wysocki S & Brzozowski AM (1998) Fluorescence study of fungal

lipase from Humicola lanuginose. J Photochem Photobiol B 45, 95-102.

26. Freier D, Mothershed CP & Wiegel J (1988) Characterization of Clostridium

thermocellum JW-20. Appl Environ Microbiol 54, 204-211.

139

27. Winkler UK & Stuckmann M (1979) Glycogen, hyaluronate, and some other

polysaccharides greatly enhance the formation of exolipase by Serratia marcescens.

J Bacteriol 138, 663-670.

28. Ausubel FM, Brent R, Kingston RE, Moore DD, Smith AJ, Seidman J & Struhl K

(1995) In Current Protocols in Molecular Biology. Greene press, NY, USA.

29. Sambrook J, Fritsch EF & Maniatis T (1989) Molecular cloning: A laboratory manual,

2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA.

30. Brito RM & Vaz, WL (1986) Determination of the critical micelle concentration of

surfactants using the fluore-scent probe N-phenyl-1-naphthylamine, Anal Biochem

152, 250-255.

31. Chirita CN, Necula M & Kuret J (2003) Anionic micelles and vesicles induce tau

fibrillization in vitro. J Biol Chem 278, 25644-25650.

140

Table 3.1. Properties of various detergents.

Detergent MW Class CMC

(mM)*a

CMC

(mM)b

CMC

(mM)c

Triton X-100 625 (avgas) Nonionic 0.24 0.26 0.24

Tween 20 1228 (avg) Nonionic 0.06 0.066 0.058

Tween 80 1310 (avg) Nonionic 0.01 0.014 0.01

CTAB 364 Cationic 0.92 1.1 0.9

Na cholate 430.5 Anionic 14.0 14.6 14.6

SDS 288 Anionic 2.0 2.2 2.0

* The error on the cmc determinations is 10%.

a Literature values [9]

b Measured in 100mM TAPS pH 9.0.

c Measured in 100 mM TAPS pH9.0 and 3mM pNPL

141

Table 3.2. The effect of E600 on LipA and LipB activity in the presence of SDS and

Tween 20.

[SDS]a LipAb

(U·mg-1)

LipBc

(U·mg-1)

LipA+E600

(U·mg-1)

LipB+E600

(U·mg-1)

0 11.5 ± 0.6 11.8 ± 0.6 8.6 ±1.0 9.0 ± 1.0

1mM 7.6 ± 1.0 9.0 ± 1.0 6.0 ± 1.0 6.2 ± 1.0

3 mM 7.8 ± 2.0 8.4 ± 2.0 2.0 ± 1.0 2.3 ± 1.0

7mM 8.0 ± 2.0 8.5 ± 2.0 3.0 ± 1.4 1.6 ± 1.4

[Tween 20]a LipA

(U·mg-1)

LipB

(U·mg-1)

LipA+E600

(U·mg-1)

LipB+E600

(U·mg-1)

0 11.6 ± 0.5 12.4 ± 0.5 9.6 ± 1.0 9.0 ± 1.0

0.008 mM 10.0 ± 1.2 9.5 ±1.6 5.0 ± 1.4 5.4 ± 1.5

0.01 mM 9.0 ± 2.0 9.2 ± 2.0 3.0 ± 2.0 1.0 ± 0.5

0.04 mM 7.6 ±1.2 7.0 ± 1.4 3.4 ± 1.6 1.8 ±1.4

0.08 mM 7.0 ± 1.0 6.8 ± 1.4 2.4 ± 1.0 3.5 ±1.6

1.0 mM 6.4 ± 1.8 6.5 ± 1.8 3.0 ± 1.4 3.8 ± 1.4

a Thermostability of LipA and LipB is greatly reduced by SDS and Tween 20.

b In the absence of detergents, LipA and LipB retained 90% activity after incubation at 60°C for 3 h.

c LipA and LipB specific activities before incubation were 12.4 and 13.0 U·mg-1, respectively.

142

Fig. 3.1.

Temperature activity profile with 0.2% SDS. Assays were performed as described in

the materials and methods section. The assays were run in triplicate at the pH optima of

LipA (○) and LipB (□). Inserted window is the temperature activity profile without SDS

expressed in U·mg-1. All data were recorded as the difference between the assays and

their controls.

143

144

Fig. 3.2.

The hydrolysis rate of p-nitrophenyl laurate by LipA in the absence (●) and

presence of SDS (○). Assays were performed as described in the materials and

methods section. Each assay contained 10 µg of freshly purified LipA. The assay

duration was 15 minutes, at the end of the assay; reactions were stopped in ice. Since

LipA and LipB look very much the same, LipB was omitted for clarity.

145

146

Fig. 3.3.

Effects of various detergents on the activity of LipA (○) and LipB (□). Increasing

concentrations of the following detergents were used; SDS (A), Na cholate (B), CTAB

(C), Triton X-100 (D), Tween 20 (E) and Tween 80 (F). Assays were carried out as

described in the materials and methods section with some modifications. The assay

buffer was 100 mM TAPS adjusted at pH25°C9.0. The assay temperature was 92 °C.

Running assays containing non ionic detergents at concentrations higher than their

CMC values cause turbidity in the solution, to overcome this problem, assays were

centrifuged and diluted.

147

148

Fig. 3.4.

The effect of detergent on stability of LipA (A) and LipB (B) at room temperature.

The following detergents were used: SDS (♦), Tween 20 (■), their effects on enzymes

stability were compared to enzyme stability in the absence of detergent (○). Assays

were carried out as described in the materials and methods section.

149

150

CHAPTER 4

OLIGOMERIZATION OF TWO THERMOPHILIC LIPASES AND ITS EFFECT ON

THERMOSTABILITY1

1Salameh, M. and J. Wiegel. 2006. To be submitted to FEMS Microbiology Letters

151

Abstract

Two thermophilic extracellular lipases, LipA and LipB, purified to gel electrophoretic

purity from Thermosyntropha lipolytica DSM 11003, were found to exhibit a strong

tendency to oligomerize as observed through gel filtration chromatography and gradient

native gel electrophoresis. The addition of 1% (w/v) SDS lead to deoligomerization as

the lipases eluted with an apparent peak corresponding to their monomeric mass based

on SDS gel electrophoresis. Both enzymes have remarkable thermostability; the effect

of oligomerization on thermostability was tested by incubating the enzymes in the

presence of ionic and non ionic detergents. The steep decrease in thermostability upon

incubation of the enzymes with ionic and non ionic detergents suggests that, enzyme

aggregation might be a major contributor to their high thermostability.

152

Introduction

Thermosyntroph lipolytica DSM 11003, a thermophilic, alkalitolerant, lipolytic, syntrophic

anaerobic bacterium was isolated in 1996 from Lake Bogoria in Kenya (Svetlitshnyi, et

al., 1996). It was isolated with the specific intent of finding bacterial lipases with

tolerance for, and activity, at high temperatures and pH values. True Lipases (E.C.

3.1.1.3) are carboxyl ester hydrolases that catalyze the hydrolysis and synthesis of long

chain acylglycerols (Sarda & Desnuelle, 1958; Ferrato et al., 1997), whereas esterases

exhibit only activity with short chain acylglycerols.

We have recently described the purification and characterization of two extracellular

lipases, LipA and LipB, from T. lipolytica (Salameh & Wiegel, submitted (a)). LipA and

LipB both exhibit maximum catalytic activity at 96 °C, which is the highest among all

published lipases, and an optimum pH25°C at 9.4 and 9.6, respectively. Both LipA and

LipB show high thermostability, they retain 50% activity after 20 hours incubation at 75

°C.

Several lipases were found to form “aggregation”, most noticeably are the lipases

from Burkholderia (bas Pseudomonas) cepacia (Dünhaupt et al., 1992), Geobacillus

(bas Bacillus) thermocatenulatus (BTL2) (Rua et al., 1997), rice bran (Oryza sativa)

(Bhardwaj et al., 2001), B. subtilis (Lesuise et al., 1993), Most interestingly the lipases

from four strains of the psychrotroph Moraxella sp. are also secreted in large

“aggregates” (~400kDa) (Feller et al., 1990). At this time, it is unclear whether the use of

the term “aggregation” is appropriate, or whether one should differentiate between

aggregation and oligomerization.

153

In thermophilic organisms, an increase in oligomerization state is one of the protein

stabilization strategies that is observed (Walden et al., 2001). Protein monomers

interface is dependent on the nature and affinity of the interactions comprising that

interface, but both hydrophobic and polar interactions play key roles in most interfaces

(Ali & Imperiali, 2005). Here, we report on the oligomerization of LipA and LipB and the

effects of detergents such as, SDS and Tween 20 on their thermostability.

Materials and Methods

Culture conditions

T. lipolytica was grown in a basal medium containing 0.75% yeast extract as carbon and

energy sources under a nitrogen gas phase as described elsewhere (Salameh and

Wiegel, submitted (a)). The pH of the medium was adjusted at 8.225°C and the growth

temperature was at 60 °C.

Lipase assay

Lipase assays were carried out as described previously (Salameh and Wiegel,

submitted) using a spectrophotometric method with p-nitrophenyl laurate (pNPL)

(Sigma). One unit is defined as the amount of the enzyme catalyzing the release of 1

µmol of p-nitrophenol per min from pNP-laurate/palmitate.

Gel Filtration

Concentrated aliquots (0.6mg/ml) of LipA and LipB were loaded onto Superose12

column (1.8 cm × 30 cm) and eluted by 20 mM Tris buffer pH 8.0 containing 100 mM

NaCl at 0.5 ml·min-1. A standard curve was constructed by running the following

molecular marker (Sigma USA) for gel filtration chromatography: blue dextran

154

2000 (2,000 kDa), bovine serum albumin (66 kDa), albumin (45 kDa), carbonic

anhydrase (29 kDa) and cytochrome c (12.4 kDa).

Gel electrophoreses

Electrophoretic runs were performed with a Bio-Rad Mini-Protean II cell unit, at room

temperature. SDS/PAGE and non denaturing PAGE were performed essentially as

described by (Sambrook et al., 1989; Ausubel et al., 1995). Gels were stained with

GelCode® Blue Stain Reagent (Pierce). The molecular mass of the denatured enzyme

was obtained by interpolation on a plot of the log of molecular mass against relative

migrations (RF values).

Protein concentration

Protein concentrations were determined with the bicinchoninic acid (BCA) protein assay

kit (Pierce) and follow the manufacturer instructions.

The effect of detergents on stability

Aliquots of purified LipA and LipB (Salameh and Wiegel, submitted) were incubated in

the presence of 1% detergent in sealed 2 ml serum bottles in triplicates. For every time

point three incubations were sacrificed. Each sample contained 15 µg of enzyme in 0.6

ml of 100 mM Tris buffer, pH 8.0 and 0.5 M ammonium sulfate. After incubation, the

samples were concentrated and desalted by using 10 kDa cut off centricon filter tubes

(Millipore), protein quantified as mentioned earlier and then assayed for lipase activity.

Results and Discussion

Oligomerization of LipA and LipB

LipA and LipB form oligomers to some extent as was observed during gel filtration

chromatography (Fig. 4.1). When aliquots of concentrated LipA and LipB were loaded

155

onto Superose12 column and eluted by using 20mM Tris buffer containing 100 mM

NaCl, LipA and LipB eluted at broad peaks corresponding to a molecular mass

corresponding to 280 kDa and 400 kDa, respectively. The high molecular weight

oligomers were also observed on a gradient native PAGE (Fig. 4.2A), where LipA and

LipB showed a migration corresponding to an estimated value of 300 kDa and 400 kDa,

respectively. Fractions which represent high molecular weight oligomers after gel

filtration were pooled, concentrated, and after determining specific activity analyzed on

SDS-PAGE (Fig. 4.2B). All these results suggested that LipA and LipB existed in

catalytically active and soluble oligomers, it also suggested that both enzymes posses a

considerable surface hydrophobicity, although it seemed that LipB display larger

aggregation and stronger hydrophobic interactions. The fact that when mixing LipA and

LipB and analyzing them on native gel, they exhibited two distinct bands identical to the

corresponding LipA and LipB. This indicated that there is no affinity to oligpmerize or

aggregate between LipA and LipB molecules (Fig. 4.2A).

Organic solvent and detergents were successfully used to disrupt the high molecular

weight aggregation of lipases (Dünhaupt et al., 1992; Rua et al., 1997; Bhardwaj et al.,

2001; Schlieben et al., 2004; Graupner et al., 1999). SDS was proven to activate LipA

and LipB by promoting open conformation (Salameh and Wiegel, submitted (b)); we

used 1% SDS to resolve the oligomers of LipA and LipB. In the presence of 1% (w/v)

SDS during gel filtration analysis, one peak corresponding to the elutional monomers of

LipA and LipB (Fig. 4.1). When instead of the activating SDS (Salameh and Wiegel,

submitted (b)) 40% (v/v) 2-propanol was used (Dünhaupt et al., 1992), which

deaggregate the lipase, a considerable loss of activity (80%) was observed.

156

The oligomerization and/or aggregation of proteins are primarily due to hydrophobic

interactions between their domains (Ali & Imperiali, 2005; Schmidt-Dannert et al., 1994).

The most noticeable feature of described lipase oligomerization/aggregation is that all

exhibited catalytic activity (Schlieben et al., 2004; Rua et al., 1997; Lesuise et al., 1993,

Feller et al., 1990). However, it needs to be noted that the lipase aggregation described

is regarded as being different from the common protein aggregation that resulted as a

consequence of conformational alterations attributed to denaturation and hence protein

inactivation (Joly, 1965; Remmele et al., 2005). It is believed that this type of protein

aggregation arises from the exposure of buried hydrophobic groups in the unfolded

state followed by nonspecific association of these groups (Remmele et al., 1999)

Protein oligomerization and thermostability

There are mixed results regarding the effect of aggregation on catalytic activity of

lipases. For example, Dünhaupt et al. reported a substantial increase in catalytic activity

of B. cepacia lipase as a result of deaggregation by the addition of 2-propanol. On the

contrary, Luisa Rúa et al. used cholate to deaggregate G. thermocatenulatus lipase

(BTL2), and found no effect on catalytic activity. In case of LipA and LipB, no effect on

catalytic activity by the oligomerization was observed, no increase or decrease in

catalytic activity was observed by treating the lipases with 0.1 to 1% (v/v) Tween 20

(Salameh and Wiegel, submitted (b)).

Oligomerization is one of the means of by which proteins can be stabilized in

thermophiles (Walden et al., 2001). Thus the thermostability of LipA and LipB were

tested in the monomeric and oligomeric form. Compiling the data from literature no

obvious relationships were found between stability at high temperature and degree of

157

aggregation. For example, B. cepacia lipase forms aggregates and exhibits high

thermostability (t1/2 13h at 90°C), (Dünhaupt et al., 1992), while G. thermocatenulatus

lipase (BTL2) aggregated form has a half life of 30 min at 60°C (Schmidt-Dannert et al.,

1994; Rua et al., 1997). However, as mentioned previously, LipA and LipB exhibited

high thermostability (LipA: t1/2 6 h at 100 °C, 20h at 75 °C. LipB: t1/2 2h at 100 °C, 22h at

75 °C at pH 8), in addition to their physical properties, two factors were described that

might contribute to this high thermostability, first, the addition of manganese ions (Mn2+ )

was found to increase thermostability, most probably, by increasing electrostatic

interactions (Salameh and Wiegel, submitted (a)). Secondly, the presence of 0.5-2 M

ammonium sulfate, which promotes hydrophobic interactions, was found to increase the

half times for inactivation of both lipases at elevated temperatures (from 30 h to 48 h at

60 °C). This suggests the possibility that as the oligomerization is hydrophobic in nature,

ammonium sulfate might promote and stabilized this oligomerization of LipA and LipB

and as a result enhances thermostability. To test this hypothesis, the thermostability of

LipA and LipB at 75 °C was analyzed in the presence of ionic detergent SDS and non

ionic detergent Tween 20. In the presence of either detergent, the lipases were in the

monomeric form as determined by gel filtration. Both detergents had a profound effect

on destabilizing the enzymes. The half lives of both enzymes were reduced by SDS and

Tween 20 to 1 and 2 hours down from 24 hours, respectively (Fig. 4.3). Nonionic

detergents normally are considered as mild detergents and that they do not interact

extensively with the protein surface, whereas ionic detergents, in particular SDS,

generally bind unspecifically to the protein surface, which usually lead to protein

158

unfolding (Mogensen et al., 2005). However, both detergents exhibited similar and

dramatic effects on thermostability.

Thermophilic proteins in general are thought to show more stability at room

temperature because they are less flexible, but as the temperature increases, proteins

become more flexible and this increases the unfolding rate and decreases enzymes

stability. SDS in contrast to nonionic detergent Tween 20, is very reactive with LipA and

LipB, enzymes became more flexible and the rate of unfolding increased. Consequently,

the sudden decrease in thermostability of LipA and LipB in the presence of 0.1% (v/v)

Tween 20 suggested that the aggregation behavior of these enzymes might play an

important role for their high thermostability.

159

References

Ali MH & Imperiali B (2005) Protein oligomerization: how and why. Bioorg Med

Chem 13:5013-20.

Ausubel FM, Brent R, Kingston RE, Moore DD, Smith AJ, Seidman J & Struhl K (1995)

In Current Protocols in Molecular Biology. Greene press, NY, USA.

Bhardwaj K, Raju A & Rajasekharan R (2001) Identification, purification, and

characterization of a thermally stable lipase from rice bran. A new member of the

(phospho) lipase family. Plant Physiol 127: 1728-1738.

Dünhaupt A, Lang S & Wagner F (1992) Pseudomonas cepacia lipase: studies on

aggregation, purification and on the cleavage of olive oil. Biotechnol Lett 14: 953–

958.

Feller, G. Thiry, M. Arpigny, J. L. Mergeay, M. and C. Gerday. 1990. Lipases from

psychrotrophic antarctic bacteria. FEMS Micriobiol. Lett. 66: 239-244.

Ferrato F, Carriere F, Sarda L & Verger R (1997) A critical reevaluation of the

phenomenon of interfacial activation. Methods Enzymol 286: 327-347.

Graupner M, Haalck L, Spener F, Lindner H, Glatter O, Paltauf F & Hermetter A (1999)

Molecular dynamics of microbial lipases as determined from their intrinsic tryptophan

fluorescenc. Biophys J 77: 493-504.

Joly M (1965) Physico-Chemical Approach to the Denaturation of Proteins; Academic

Press, London, UK.

Lesuise E, Schanck K & Colson C (1993) Purification and preliminary characterization

of the extracellular lipase of Bacillus subtilis 168, an extremely basic pH-tolerant

enzyme. Eur J Biochem 216: 155–160.

160

Mogensen JE, Sehgal P & Otzen DE (2005) Activation, Inhibition, and Destabilization of

Thermomyces lanuginosus Lipase by Detergents. Biochemistry 44: 1719 -1730.

Remmele Jr. RL, Bhat SD, Phan DH & Gombotz WR (1999) Minimization of

recombinant human Flt3 ligand aggregation at the T plateau: A matter of thermal

reversibility.

m

Biochemistry 38: 5241-5247.

Remmele Jr. RL, Zhang-van Enk J, Dharmavaram V, Balaban D, Durst M,

Shoshitaishvili A & Rand H (2005) Scan-rate-dependent melting transitions of

interleukin-1 receptor (type II): elucidation of meaningful thermodynamic and kinetic

parameters of aggregation acquired from DSC simulations. J Am Chem Soc 127:

8328–8339.

Rua ML, Schmidt-Dannert C, Wahl S, Sprauer A & Schmid RD (1997)

Thermoalkalophilic lipase of Bacillus thermocatenulatus large-scale production,

purification and properties: aggregation behaviour and its effect on activity. J

Biotechnol 56: 89-102.

Salameh M & Wiegel J (2006 a) Purification and characterization of two highly

Thermophilic alkaline lipases from Thermosyntropha lipolytica. Appl Environ

Microbiol (submitted).

Salameh M & Wiegel J (2006 b) Effects of various detergents on two alkalithermophilic

lipases from Thermosyntropha lipolytica. FEBS J (submitted).

Sambrook J, Fritsch EF & Maniatis T (1989) Molecular cloning: A laboratory manual,

2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA.

Sarda L & Desnuelle P (1958) Action of pancreatic lipase on emulsified esters. Biochim

Biophys Acta 30: 513-521.

161

Schlieben NH, Niefind K & Schomburg D (2004) Expression, purification, and

aggregation studies of His-tagged thermoalkalophilic lipase from Bacillus

thermocatenulatus. Protein Expr Purif 34:103-110.

Schmidt-Dannert C, Sztajer H, Stocklein W, Menge U & Schmid, RD (1994) Screening,

purification and properties of a thermophilic lipase from Bacillus thermocatenulatus.

Biochim Biophys Acta 1214: 43-53.

Svetlitshnyi V, Rainey F & Wiegel J (1996) Thermosyntropha lipolytica gen. nov., sp.

nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing short and

long chain fatty acids in syntrophic coculture with a methanogenic archaeum. Int J

Syst Bacteriol 46:1131-1137.

Walden H, Bell GS, Russell RJM, Siebers B, Hensel R & Taylor GL (2001) J Mol Biol

306:745–757.

162

Fig. 4.1.

The elution profiles of aggregated LipA (a) and LipB (b) after gel filtration

chromatography. The first broad peaks represent aggregated enzymes, the second

sharp peaks represent enzymes treated and eluted with 1% SDS. The elution volume

and corresponding molecular mass as calculated from standard marker is indicated on

top of each peak. Elution volumes of marker proteins were as follows: blue dextran

(2000 kDa), 21.90 ml, bovine serum albumin (66 kda), 33.56 ml, albumin (45 kDa),

36.47 ml, carbonic anhydrase (29kDa), 39.94 ml, cytochrome (12.8 kDa), 41.67 ml.

163

164

Fig. 4.2.

(A) 12% SDS-PAGE of eluted fractions after gel filtration of aggregated LipA. (B)

16% Gradient native PAGE of purified LipA and LipB. Lane 1, LipA, lane 2, mixed

LipA and LipB, lane 3, molecular marker.

165

166

Fig. 4.3.

Thermostability profiles of LipA (A) and LipB (B) at 75 °C. A15µg aliquots of LipA

and LipB were stored in the presence of 1% (v/v) Tween 20 (■) and 1% (w/v) SDS (♦),

in addition to the control without detergent (○). Lipase assays were conducted as

mentioned in the methods section.

167

168

CHAPTER 5

SYNTHESIS OF FATTY ACID ESTERS AND DIACYLGLYCEROLS AT ELEVATED

TEMPERATURES BY ALKALITHERMOPHILIC LIPASES1

1Salameh, M. and J. Wiegel. 2006. To be submitted to Journal of Applied and Industrial Microbiology

169

Abstract

LipA and LipB of Thermosyntropha lipolytica DSM 11003 are the most alkalithermophilic

and thermostable lipases currently known. The partially purified enzymes were

analyzed for their ability to catalyze synthesis of diacylglycerols and various alcohol fatty

acids in organic solvents. Lyophilization of LipA and LipB resulted in 40% and 50% loss

of catalytic activity, respectively, and this was overcome by the addition of 2 mg/ml of

bovine serum albumin (BSA) and 25% polyethylene glycol (PEG), resulting in a

recovery of 100% activity. Isooctane was found to be the most efficient solvent for

esterification reactions at 85 °C. The highest yield in esterification using fatty acids and

alcohols was 25% conversion using octyl oleate (LipA) and lauryl oleate (LipB). In

addition, LipA and LipB catalyzed the synthesis of 1,3-dioleoyl glycerol, 1- oleoyl-3-

lauryl glycerol and 1-oleoyl-3-octoyl glycerol. The synthesis of diglycerides was

achieved by using 1-oleoyl glycerol as substrate; substituting it with 1-lauroyl glycerol

resulted in poor diglycerides synthesis. Similar to the positional specificity in aqueous

solutions, LipA and LipB catalyzed the synthesis of diacylglycerol at 1 and 3 positions,

which make them potentially valuable for industrial applications particularly in structured

lipid biosynthesis.

170

Introduction

Thermosyntroph lipolytica, a thermophilic, alkalitolerant, lipolytic, syntrophic anaerobic

bacterium was isolated with the primary intent of finding a commercially viable lipase for

use in high temperature, alkaline environments such as those used in laundering

clothes. The organism grew on triacylglycerols and ferment saturated and unsaturated

fatty acids but not the glycerol [35]. Lipases (carboxyl ester hydrolases, E.C. 3.1.1.3)

are enzymes that generally catalyze the synthesis and hydrolysis of long chain fatty acid

esters [13, 33] (Fig. 1a). They are ubiquitous in nature, produced by both higher and

lower eukaryotes as well as many bacteria, but they have not yet been found in archaea.

T. lipolytica constitutively produces two lipases, LipA and LipB. They are the most

thermophilic lipases reported, so far, with maximum catalytic activity at 96 °C (20 min

assay), and optimum pH80°C around 8.5. They show high thermostability, retaining 50%

of their activity after a 24 hour incubation at 75 °C, and have maximum catalytic activity

with glycerides containing long chain fatty acids [31].

Lipases are increasingly used to catalyze enantioselective reactions for the

synthesis of fine chemicals and the kinetic resolution of racemates [28]. In addition, they

are used in laundry detergent, biodiesel production, lubricants, cosmetic formulations

and flavor and aroma constituents [5, 6, 17-19, 34, 36].

Thermophilic enzymes are generally more resistant than mesophilic ones to

denaturation in organic solvents [10]. The low solubility of substrates in organic solvents

is a major obstacle [3, 38] that could be avoided by conducting esterefication reactions

at elevated temperatures. The demand for fatty acids has been growing by

approximately 4% per year to reach 3,000,000 metric tons per year. The natural fatty

171

acids are mainly obtained from the hydrolysis of hard animal fats (tallow), coconut, palm

kernel and soybean oils. Almost 100 thousand tons of fatty acids are consumed to

prepare various fatty acid esters as estimated by Rohm and Haas Company

(www.rohmhaas.com). A wide variety of fatty alcohols are used in manufacturing soaps

and detergents, cosmetics, wood preservatives and personal care products

(www.icislor.com). Various fatty alcohols are also produced as biodegradable

replacements for mineral oil as textile lubricants in spin finishes, to enhance the gloss in

stick make-up and hair grooming products, used as an emollient for all types of creams

and lotions and a lubricant for plastics and metal working industries

(www.thornleycompany.com).

Lipases are used to synthesize diacylglycerols (DAG), which have multifunctional

and nutritional properties. They are constituents of edible fats and oil. For example, a

diet containing DAG and, especially, sn-1,3-diacylglycerols were found to reduce total

fat content in men and reduce obesity [25, 26]. They were also used successfully as

part of a reduced-energy diet that enhances loss of body weight, despite having similar

energy value and digestibility to triacylglycerols (TAG) [23, 40]. As a result, 1,3-

diacylglycerols were introduced in Japan as cooking oil under the trade name of Econa

to reduce body fat accumulation [22].

Previously, we reported on the purification and characterization of two highly

thermophilic lipases, LipA and LipB. Here, we report on the ability of these two lipases

to catalyze in organic solvents the esterification reactions of various alcohols with oleic

acid at elevated temperatures. This is the first report to describe esterification reaction

catalyzed by lipases at temperatures as high as 90 °C.

172

Materials and Methods

Culture conditions

T. lipolytica was grown in a basal medium containing 0.75% yeast extract as carbon and

energy sources under nitrogen gas phase as described elsewhere [31]. The pH of the

medium was adjusted at 8.225°C and the growth temperature was 60 °C.

Lipase assay

Lipase was assayed by two methods. The first method used the chromogenic substrate

p-nitrophenyl laurate (pNPL) (Sigma USA) [31]. One unit was defined as the amount of

the enzyme catalyzing the release of 1 µmol of p-nitrophenol per min from pNPL. The

second method measured the formation of unesterified free fatty acids using the NEFA

C kit (Waco USA) according to the manufacture’s instructions.

Partial purification of lipases for synthesis in organic solutions

After 18 h of growth of T. lipolytica, cells were removed from the culture broth by

collection on an Amicon hollow fiber with one million cut off. The proteins in the

supernatant were then concentrated on a 10 kDa Amicon hollow fiber (Millipore)

precipitated by the slow addition of four volumes of cold (-20 °C) acetone. The white

precipitate was collected by centrifugation and then dissolved in a 20 mM TAPS buffer,

pH 9.0. The protein solution was then loaded onto an Octyl Sepharose fast flow column

(Amersham Biosciences) (30 ml bed volume), and the two lipases, LipA and LipB, were

eluted separately at 1.8 M and 0.5 M ammonium sulfate in Tris buffer pH 8.0,

respectively [31].

Gel electrophoreses

173

Electrophoretic analyses were performed with a Bio-Rad Mini-Protean II cell unit, at

room temperature. SDS/PAGE and non denaturing PAGE were performed essentially

as described by Ausubel [4] and Sambrook [32]. Gels were stained with GelCode® Blue

Stain Reagent (Pierce).

Protein concentration

Protein concentrations were determined with the bicinchoninic acid (BCA) protein assay

kit (Pierce) and follow the instructor’s directions.

Enzyme lyophilization

Aliquots of LipA (50 µg) and LipB (100 µg) were equilibrated with 100 mM TAPS buffer

pH 9.0 containing 1mM MnCl2 [31], 25% polyethylene glycol (v/v) and 2 mg/ml bovine

serum albumin (BSA). Samples (1 ml) of the enzymes were placed into 10 ml serum

bottles, freezed and then lyophilized in a freeze dryer.

Synthesis of fatty acid alcohol esters

The esterification mixtures were composed of 100 mM of fatty acids and 100 mM of

alcohols in 2 ml of isooctane. Reactions were conducted at 85 °C with a shaking speed

at 300 rpm (Fig. 1b). Quantitative analysis was done by measuring the unesterified free

fatty acids by using NEFA C kit. Qualitative analysis was done by thin layer

chromatography.

Synthesis of diacylglycerols (DAG)

Synthesis of DAG were conducted in 10 ml serum bottles that contain 100 µg of

lyophilized LipA and LipB, 100 mM monoacylglycerols and 100 mM of fatty acids at 85

°C for 100 hours. The shaking speed was 300 rpm. The unesterified free fatty acids

174

were measured using the NEFA C kit. Qualitative analysis was done by thin layer

chromatography as described below.

Effect of temperature on activity and stability in isooctane

The effect of temperature on activity was investigated by determining the esterification

of 1-heptanol, 1-octanol and 1-dodecanol and oleic acid in isooctane. The esterification

reaction was carried out in isooctane (3 ml) containing 50 mM of alcohol and 50 mM of

oleic acid with 50 µg of lyophilized enzyme. The vials were incubated at various

temperatures with shaking at 300 rpm for 60 hours. Samples were withdrawn for TLC

analyses. For quantitative measurements, samples were withdrawn, concentrated, and

then used to quantify the non esterified fatty acids (NEFA) by using NEFA C kit. To

estimate LipA and LipB stability in isooctane, 100 µg of lyophilized LipA and LipB were

suspended in 1 ml of isooctane in sealed serum bottles and incubated at 85 °C, 100 µl

aliquots were withdrawn and assayed for enzymatic activities in TAPS buffer.

Thin layer chromatography (TLC)

Aliquots were withdrawn from the reaction mixture to qualitatively and semiquantitatively

analyze the synthesis of DAG and fatty acids alcohol esters (butyl oleate, heptyl oleate,

octyl oleate, nonanyl oleate and lauryl oleate) by thin layer chromatography. The

aliquots were concentrated by evaporation and then applied on reversed phase (Multi-K)

TLC plates (Whatman USA). The plates were developed first with a mixture of

petrolum:diethylether:acetic acid (70:30:1) while the solvent front reached half of the

plate, then continued with chloroform:acetone: acetic acid (96:4:1). Once the solvent

has reached the end of the plate, they were dried and spots were visualized by spraying

the plates with iodine vapor (0.1% iodine in chloroform).

175

Results and Discussion

Lipases purification and lyophilization

The two extra cellular lipases LipA and LipB were partially purified from T. lipolytica by

acetone precipitation and hydrophobic interaction chromatography. The purification

yield was 62% for both enzymes combined with a purification factor of 50, and a final

specific activity of 12.0 U·mg-1 for LipA and 12.8 U·mg-1 for LipB. Enzymes were

adjusted at the pH optimum. Enzymes have pH memory, that is their ionization groups

and thus their catalytic activity in organic solvents is a reflection to the pH of the last

aqueous solution to which they were exposed [43]. As a result, their catalytic activity

can be maximized if enzymes are lyophilized from aqueous solutions at their optimal pH.

The lyophilization caused 40% and 50% inactivation of LipA and LipB, respectively

[16]. However, when 2 mg/ml of bovine serum albumin (BSA) and 25% polyethylene

glycol (PEG) (v/v) were added to the enzyme solution prior to lyophilization as structure-

preserving lyoprotectant [11], full recovery of enzyme activity and 5 times higher

esterification rates were achieved (Fig. 5.2). When protein solutions were lyophilized

after acetone precipitation, no loss of activity was detected. This could be due to the

presence of proteins and lipids in the culture supernatant that kept the enzymes in their

native, enzymatically active conformations [12]. PEG was not used by any of the lipases

as an acyl acceptor.

Choice of organic solvents

Isooctane was the most effective solvent to conduct synthesis of fatty acid alcohol

esters. After 60 hours, LipA catalyzed the conversion of 25% and 21% of octyl oleate

and lauryl oleate, respectively. LipB showed similar efficiency in isooctane, however,

176

lauryl oleate synthesis was slightly favored (Table 5.1). The choice of the organic

solvent is known to have a profound impact on esterefication reactions. Enzymes are

generally more thermostable in hydrophobic solvents than hydrophilic ones [37], the

type of organic solvent affects the conversion rates of esterification and condensation

reactions [1, 9, 30, 39] and enantioselectivity [8, 14, 24, 27, 41, 42].

Acetonitrile, was not an effective solvent with which to conduct synthesis of fatty acid

alcohol esters (Table 5.1). Water-immiscible organic solvents such as isooctane are

inert with respect to their interaction with proteins; miscible solvents such as acetonitrile

tend to strip water molecules from the enzyme which are essential for catalytic activity

[37]. In general, enzymes have much lower catalytic activity in organic solvents, since

they lack structural flexibility [21, 29]. In aqueous environments, enzymes are more

flexible, they form hydrogen bonds with water molecules, the lack of these hydrogen

bonds in organic sovent lead to stronger intra-protein electrostatic interactions [2].

Consequently, enzyme molecules become much more rigid [7, 20].

Temperature effect and fatty alcohols synthesis

The maximum temperature for catalytic activity in organic solvents was determined by

conducting the esterification reactions in isooctane at different temperatures. LipA and

LipB have a maximum activity range at 85 °C to 90 °C (Fig. 5.3a). Regarding

thermostability, i.e. the gradual, irreversible loss of enzymatic activity upon exposure to

high temperatures, an enzyme could have a short half-life in aqueous solution, however

it might exhibit high stability in organic solvents. For example, porcine pancreatic lipase

withstands heating at 100 °C for many hours in an organic solvent [44]. Likewise, the

thermal denaturation of a ribonuclease was greatly enhanced in organic solvent (Tm

177

values as high as 124 °C) compared to aqueous solution (61 °C Tm value) [37]. In this

study we show that both lipases lose about 35% of their activity after 24 hours of

incubation, with half lives near 48 hours. However, 25% of residual activity was detected

after 96 hours of incubation (Fig. 5.3b). The decrease in thermal stability might be

attributed to the presence of small water activity in the lyophilized enzymes preparation.

The presence of extremely low water along with the organic solvent has a dramatic

effect on lowering stability of proteins [15].

As shown in Figure 5.4, both enzymes catalyze the esterification of oleic acid with all

middle and long chain alcohols tested, including 1-heptanol, 1-octanol, 1-nonanol and 1-

dodecanol, but not with 1-butanol. The highest conversion was achieved by LipA

yielded 25% octyl oleate and 23% lauryl oleate. The maximum conversion by LipB was

24% of lauryl oleate and 21% of octyl oleate. Qualitative measurements of lipase-

catalyzed fatty alcohols synthesis were monitored by TLC through out the reaction time

course (Fig. 5.5).

Both LipA and LipB were proven to show maximum hydrolysis of lipids with long

chain unsaturated fatty acids and to lesser extent saturated long chain fatty acids but

not short or middle range fatty acids [31]. Indeed, oleic acid was the preferred acyl

donor in esterefication reactions; the results were quite different once oleic acid was

substituted with palmitic acid, lauric acid and butyric acid as acyl donor. Only 8%

conversion of octyl palmitate and octyl laurate and 0% of octyl butyrate occured after 60

hours. This leads to the conclusion that short chain acyl donors or acceptors are not

preferred substrates for LipA and LipB. The low conversion rate observed could be

178

attributed to the low concentration of the starting enzymes and the low catalytic activity

and stability.

Diacylglycerols synthesis

Diacylglycerols (DAG) were synthesized by LipA and LipB-catalyzed esterification

reactions, using 1-olyel acylglycerol and 1-lauryl acylglycerol and various fatty acids as

substrates (Fig. 7). It is evident from the data provided that both lipases lead to

extensive incorporation of unsaturated long chain fatty acid (e.g. oleic acid) into

diacylglecerol. The results also demonstrate that 1,3-dioleylglycerol has the highest

conversion percentage (62%) among all of the DAGs generated. In contrast, the

synthesis of DAG from 1-laurylacylglycerol and various fatty acids was very low. TLC

analysis revealed no indication of any triacylglycerol synthesis which parallels the 1, 3

positional specificity of LipA and LipB during hydrolysis in aqueous solutions (31).

In conclusion, the positional specificity of LipA and LipB make them attractive

enzymes for several applications, especially for structured lipid, flavor and aroma

constituents’ synthesis where alkaline pH and elevated temperatures are used.

179

References

1. Adachi S, Kobayashi T (2005) Synthesis of esters by immobilized-lipase-catalyzed

condensation reaction of sugars and fatty acids in water-miscible organic solvent. J

Biosci Bioeng 99:87-94

2. Affleck R, Haynes CA, Clark DS (1992) Solvent dielectric effects on protein

dynamics. Proc Natl Acad Sci USA 89:5167-5170

3. Akimoto T, Nagase Y. (2003) Novel transdermal drug penetration enhancer:

synthesis and enhancing effect of alkyldisiloxane compounds containing

glucopyranosyl group. J Control Release 88:243-252.

4. Ausubel FM, Brent R, Kingston RE, Moore DD, Smith AJ, Seidman J, Struhl K

(1995) In Current Protocols in Molecular Biology. Greene press, NY, USA

5. Balcão VM, Paiva AL, Malcata FX (1996) Bioreactors with immobilized lipases:

State of the art. Enzyme Microb Technol 18:392–416

6. Bornscheuer UT, Bessler C, Srinivas R, Krishna SH (2002) Optimizing lipases and

related enzymes for efficient application. Trends Biotechnol 20:433-437

7. Burke PA, Griffin RG, Klibanov AM (1993) Solid-state nuclear magnetic resonance

investigation of solvent dependence of tyrosyl ring motion in an enzyme.

Biotechnol Bioeng 42:87-94

8. Carrea G, Ottolina G, Riva S (1995) Role of solvents in the control of enzyme

selectivity in organic media. Trends Biotechnol 13:63-70

9. Castillo E, Pezzotti F, Navarro A, lopez-Munguia A (2003) Lipase-catalyzed

synthesis of xylitol monoesters:solvent engineering approach. J Biotechnol

102:251-259

180

10. Cowan DA (1997) Thermophilic proteins: stability and function in aqueous and

organic solvents. Comp Biochem Physiol A Physiol 118:429-438

11. Dabulis K, Klibanov AM (1993) Dramatic enhancement of enzymatic activity in

organic solvents by lyoprotectants. Biotechnol Bioeng 41:566-571

12. Estrada-Mondaca S, Fournier D (1998) Stabilization of recombinant Drosophila

acetylcholine esterase. Prot Exp Purif 12:166–172

13. Ferrato F, Carriere F, Sarda L, Verger R (1997) A critical reevaluation of the

phenomenon of interfacial activation. Methods Enzymol. 286:327-347

14. Ghanem A, Schurig V (2001) Lipase-catalyzed irreversible transesterification of 1-

(2-furyl)ethanol using isopropenyl acetate. Chirality 13:118-23

15. Griebenow K, Klibanov AM, (1996) On protein denaturation in aqueous-organic

mixtures but not in pure organic solvents. J Am Chem Soc 47:11695-11700

16. Griebenow K, Klibanov AM (1995) Lyophilization-induced changes in the

secondary structure of proteins. Proc Natl Acad Sci USA 92:10969-10976

17. Jaeger K-E, Ransac S, Dijkstra B. W, Colson C, Van Heuvel M, Misset O (1994)

Bacterial lipases. FEMS Microbiol Rev 15:29-63

18. Jaeger K-E, Dijkstra BW, Reetz MT (1999) Bacterial biocatalysts: molecular

biology, three-dimensional structures, and biotechnological applications of lipases.

Annu Rev Microbiol 53:315–351

19. Jaeger K-E, Reetz MT (1998) Microbial lipases form versatile tools for

biotechnology. Trends Biotechnol 16:396-403

20. Jeffrey JA, Saenger W (1994) Hydrogen Bonding in Biological Structures. Springer

press, Berlin

181

21. Kuntz ID, Kauzmann W (1974) Hydration of proteins and polypeptides. Adv Protein

Chem 28: 239-345

22. Liao HF, Tsai WC, Chang SW, Shieh CJ (2003) Application of solvent engineering

to optimize lipase-catalyzed 1,3-diglyacylcerols by mixture response surface

methodology. Biotechnol Lett 25:1857-1861

23. Maki KC, Davidson MH, Tsushima R, Matsuo N, Tokimitsu I, Umporowicz DM,

Dicklin MR, Foster GS, Ingram KA, Anderson BD, Frost SD, Bell M (2002)

Consumption of diacylglycerol oil as part of a reduced-energy diet enhances loss

of body weight and fat in comparison with consumption of a triacylglycerol control

oil. Am J Clin Nutr 76:1230-1236

24. Miyazawa T, Kurita S, Shimaoka M, Ueji S, Yamada T (1999) Resolution of

racemic carboxylic acids via the lipase-catalyzed irreversible transesterification of

vinyl esters. Chirality 11:554-60

25. Murase T, Mizuno T, Omachi T, Onizawa K, Komine Y, Kondo H, Hase T,

Tokimitsu I (2001) Dietary diacylglycerol suppresses high fat and high sucrose

diet-induced body fat accumulation in C57BL/6J mice. J Lipid Res 42:372-378

26. Nagao T, Watanabe H, Goto, N, Onizawa K, Taguchi H, Matsuo N, Yasukawa T,

Tsushima R, Shimasaki H, Itakura H (2000) Dietary diacylglycerol suppresses

accumulation of body fat compared to triacylglycerol in men in a double-blind

controlled trial. J Nutr 130:792-797

27. Park HJ, Choi WJ, Huh EC, Lee EY, Choi CY (1999) Production of optically active

ketoprofen by direct enzymatic esterification. J Biosci Bioeng 87:545-547

182

28. Patel RN (2000) Microbial/enzymatic synthesis of chiral drug intermediates. Adv

Appl Microbiol 47:33-78

29. Rupley JA. Careri G (1991) Protein hydration and function. Adv Protein Chem

41:37-172

30. Sabally K, Karboune S, Yeboah FK, Kermasha S (2005) Lipase-catalyzed

esterification of selected phenolic acids with linolenyl alcohols in organic solvent

media. Appl Biochem Biotechnol 127:17-27.

31. Salameh M, Wiegel J, (2006) Purification and characterization of two highly

thermophilic alkaline lipases from Thermosyntropha lipolytica. Appl Environ

Microbiol (submitted)

32. Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning: A laboratory manual,

2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA

33. Sarda L, Desnuelle P (1958) Action of pancreatic lipase on emulsified esters.

Biochim Biophys Acta 30:513-521

34. Schmidt-Dannert C (1999) Recombinant microbial lipases for biotechnological

applications. Bioorg. Med. Chem. 7:2123-30

35. Svetlitshnyi V, Rainey F, Wiegel J (1996) Thermosyntropha lipolytica gen. nov., sp.

nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing short and

long chain fatty acids in syntrophic coculture with a methanogenic archaeum. Int. J

Syst Bacteriol 46:1131-1137

36. Villeneuve P, Muderhwa JM, Graille J, Haas MJ (2000) Customizing lipases for

biocatalysis: a survey of chemical, physical and molecular biological approaches. J

Mol Catal B Enzym 9:113–148

183

184

37. Volkin DB, Staubli A, Langer R, Klibanov AM (1991) Enzyme thermoinactivation in

anhydrous organic solvents. Biotechnol Bioeng 37:843-853

38. Wang N, Liu BK, Wu Q, Wang JL, Lin XF (2005) Regioselective enzymatic

synthesis of non-steroidal anti-inflammatory drugs containing glucose in organic

media. Biotechnol Lett 27:789-792.

39. Watanabe Y, Miyawaki Y, Adachi S, Nakanishi K, Matsuno R (2001) Equilibrium

constant for lipase-catalyzed condensation of mannose and lauric acid in water-

miscible organic solvents. Enzyme Microb Technol 29:494-498

40. Weber N, Mukherjee KD (2004) Solvent-free Lipase-Catalyzed Preparation of

Diacylglycerols. J Agric Food Chem 52:5347-5353

41. Wescott CR, Klibanov AM (1994) The solvent dependence of enzyme specificity.

Biochim Biophys Acta 1206:1-9

42. Xu YL, Liu ZS, Wang HF, Yan C, Gao RY (2005) Chiral recognition ability of an

(S)-naproxen- imprinted monolith by capillary electrochromatography.

Electrophoresis 26:804-11

43. Zaks A, Klibanove AM (1988) Enzymatic catalysis in non-aqueous solvents. J Bio

Chem 263:3194-3201

44. Zaks A, Klibanov AM (1984) Enzymatic catalysis in organic media at 100°C.

Science 224: 1249-1251

185

Lipase Solvent ReactionTemp. °C

Octyl oleateb

Conversion (%)

Lauryl oleate Conversion (%)

Octyl oleatec

Conversion (%)

Laury oleate Conversion (%)

LipA Isooctane 85 14 8 25 21 Octane 85 11 6 22 20 Toluene 85 6 3 11 10 Acetonitrile 75 1 2 1 1LipB Isooctane 85 12 9 23 24 Octane 85 12 8 22 21 Toluene 85 10 8 15 18 Acetonitrile 75 0 0 2 0SNa Isooctane 85 19 18 34 30 Octane 85 20 18 37 32 Toluene 85 10 8 14 11 Acetonitrile 75 3 2 5 5

Table 5.1. Lipase-catalyzed esterification of fatty alcohols in various organic solvents.

a SN is supernatant concentrate after acetone precipitation. b Reaction time was 12 h. c Reaction time was 60 h.

Fig 5.1.

The lipase reaction catalyzing the hydrolysis and synthesis of lipids (a). The

synthesis of octyl oleate (b).

186

187

Fig. 5.2.

TLC analysis of the effect of polyethylene glycol (PEG) on lipase-catalyzed octyl

oleate synthesis. Esterification synthesis and TLC analysis were carried out as

described in the methods section. The reactions were conducted for 60 hours at 85 °C.

Lane 1, Lyophilized LipA without PEG; lane 2, LipA with 25% PEG; lane 3, LipB with

25% PEG; lane 4, Control (no enzyme).

188

189

Fig. 5.3.

The effect of temperature on LipA (○) and LipB (□) activity (a) and stability (b) in

isooctane. Aliquots of enzyme solutions (100 µl) were withdrawn at various times and

used for activity assays. An enzyme sample that is suspended in isooctane without

being heated was used as reference. A 100% relative activity is 3.6 and 3.3 U·mg-1 for

LipA and LipB, respectively. Assays were carried out in triplicate

190

191

Fig. 5. 4.

Time course of fatty acid alcohol esters synthesis catalyzed by LipA (a) and LipB

(b). Butyl oleate (◊), heptyl oleate (□), octyl oleate (○), nonanyl oleate (∆) and lauryl

oleate (×). The inside windows represent TLC analysis of fatty alcohols production as

described in the methods section. Lanes from left to right are lauryl oleate, nonanyl

oleate, octyl oleate, heptyl oleate, butyl oleate and control.

192

193

Fig. 5.5.

TLC analysis and time course formation of octyl oleate (A, B) and lauryl oleate (C,

D) by LipA (A, C) and LipB (B, D).

194

195

Fig. 5.6.

Time course of diacylglycerol synthesis catalyzed by LipA (a) and LipB (b). 1,3-

Dioleoyl glycerol (◊), 1-oleoyl-3-lauroyl glycerol (□), 1-oleoyl-3-octoyl glycerol (∆), 1,3-

dilauroyl glycerol (×) and 1-lauroyl-3-oleoyl glycerol (○). TLC analysis of DAG (c). Lane1;

control (no enzyme), lane 2; 1,3-dioleoyl glycerol, lane 3; 1-oleoyl-3-octoyl glycerol, lane

4; 1-oleoyl-3-lauroyl glycerol, lane5; 1, 3-dioleoyl glycerol marker, lane 6; trioleate

marker

196

197

CHAPTER 6

CONCLUSIONS

Extremophiles includes organisms able to thrive at extremes of temperature,

pressure, low water activity, salinity, acidity, alkalinity, radiation and cmpination thereof.

As a result, extremophiles have the potential to produce uniquely valuable biocatalysts

that function under conditions in which, their usually non-extremophilic counterparts

could not.

The attempts to discover novel enzymes from extreme environment have drastically

increased in the past decade. Such enzymes serve as excellent models for

understanding protein stability and carry significant potential for biotechnology. The

most thoroughly investigated extremophilic enzymes are the thermophilic ones.

Although there are some enzymes from mesophilic sources that are found to withstand

elevated temperatures, such cases are rare. The applied interest in thermophilic

enzymes is related to their overall inherent stability which is important to performing

many biotechnologically related processes.

Thermosyntropha lipolytica produces two lipases, LipA and LipB. They are the most

thermophilic lipases reported so far with maximum catalytic activity observed at 96 °C,

and optimum pH at 9.4 and 9.6 for LipA and LipB, respectively. They are among the

most thermostable lipases; LipA showed similar stability to LipB and both retained 50%

activity after 1, 20, and 30 hours incubation at 100 °C, 75 °C and 60 °C, respectively.

The most profound effect on thermostability was observed with 0.5 to 2 M of ammonium

198

sulfate. This extended the half-lives of LipA and LipB at all temperatures, and most

noticeably was at 60°C, where the half life was 48 h for both enzymes. Both enzymes

preferred glycerides with long chain fatty acids, with maximum activity exhibited toward

trioleate (C18:1). Among the p-nitrophynyl (pNP) esters tested, pNP laurate was the best

substrate. LipA and LipB are specific enzyme which prefer hydrolysis of ester bonds at

1 and 3 positions. Metal analysis indicated that both LipA and LipB contain one ion of

Ca+2 and one ion of Mn+2 per monomeric unit. The presence of 1mM MnCl2 enhanced

the activity of both lipases by three fold and increased their thermal stability. The activity

of both lipases was inhibited by 10mM PMSF and EDTA. LipA and LipB were found to

be cold labile, irreversibly inactivated when frozen. Most of the catalytic activity of LipA

and LipB were preserved (75% and 90%, respectively), when they were frozen in

solution containing 40% glycerol (v/v) and 2 mg/ml bovine serum albumin.

To elucidate the effect of detergents on LipA and LipB activity, we chose detergents

with different head groups, alkyl chains and charge. LipA and LipB are true lipases that

are interfacially activated. SDS has the greatest impact on activity. The vmax values

increased from approximately 13.0 U·mg-1 to 105 U·mg-1 and 112 U·mg-1 and K0.5 values

decreased from 1.8 mM and 1.65 mM to 800 µM and 740 µM for LipA and LipB,

respectively. Maximum activation of enzymes by SDS occurred at the micelle level,

whereas with non ionic detergent occurred at the monomer level. SDS micelles and

Tween 20 monomers bind to the lid and activate the enzyme by triggering

conformational changes. Detergent binding cannot be identical to substrate binding;

otherwise only inhibition would be observed, not activation.

199

Both LipA and LipB have a strong tendency to form active and soluble oligomerization

of approximately 300 and 400kDa, respectively. 1% SDS or Tween 20 were able to de-

oligomerize the enzymes. The half lives of both enzymes were reduced to 1 and 2 hours

down from 22 hours by SDS and Tween 20, respectively. Enzymes oligomerization

might be a major contributor to their high thermostability.

The ability of LipA and LipB to catalyze organic synthesis was investigated. Both

enzymes lost 50% of their activity after lyophilization but this was obviated by the

addition of 25% PEG and 2mg/ml BSA. LipA and LipB have a maximum activity range

at 85°C to 90°C to conduct esterefication reactions. Isooctane as water-immiscible

organic solvent was the most effective solvent to conduct esterification reactions. LipA

and LipB showed maximum activity in the synthesis of octyl oleate and lauryloleate,

respectively. Short chain acyl donors or acceptors are not preferred substrates, 1,3-

dioleylglycerol has the highest conversion percentage at 62% among all DAGs

investigated. TLC analysis showed no indication of any triacylglycerol synthesis which

confirms the 1, 3 positional specificity of the two enzymes.

Based on all these characteristics of LipA and LipB, they can be used to conduct

several industrial applications including:

1. The production of fatty acids at elevated temperatures from beef tallow.

2. As detergent additives based on their high thermostability and thermoactivity.

3. The production of diacylglycerols.

4. The production of various fatty acid esters at elevated temperature which might

provide a solution for the low solubility of substrates in organic solvents.

200

APPENDIX A

SUPPLEMENTAL FIGURES AND TABLES

201

Table A.1. Metal analysis of purified LipA and LipB by Inductively Coupled Plasma-Emission Spectrometry.

Metal LipAa ppm

Control# ppm

∆ LipAb ppm

Ions perLipA

LipB ppm

Control ppm

∆ LipB ppm

Ions per LipB

B_2496 -0.0031 0 0 <0.05 -.0102 -.0024 0

Ba4934 0.08051 0.08105 0 <0.05 0.06110 0.08103 0

Ca3158 0.1468 0.0682 0.0786 0.92 0.1406 0.0695 0.0711 1.01

Co2286 0.00012 0.0001 0.00002 <0.05 0 0 0

Fe2599 0.0048 0.0038 0.001 <0.05 0.0037 0.0028 0.0009

K_7664 0.5669 0.6106 0 <0.05 0.5899 0.6068 0

Mg2790 0.022 0.032 0 <0.05 0.034 0.033 0.001

Mn2576 0.0636 0.0029 0.0607 0.54 0.0619 0.0044 0.00575 0.6

Ni2316 0.0044 0.0041 0.0003 <0.05 0.0036 0.004 0

Zn2139 0.0209 0.0199 0.001 <0.05 0.02340 0.0226 0.0008

a the results are based on 100µg/ml of LipA and LipB. # LipA and LipB were dialyzed against dH2O b the controls are the dialyzed buffers

202

Fig. A.1.

The time course of p- nitrophenyl laurate hydrolysis by LipA (A) and LipB (B). The

enzyme assays were conducted as described earlier (see chapter 2).The assay started

by adding 15 µg of purified enzymes. At the end of every assay, the reactions were

stopped on ice. Cleavage of pNPL was measured spectrophotometrically at 405 nm.

Average specific activity was ~13 U·mg-1.

203

204

Fig. A.2.

The catalytic activity versus enzyme concentration. The activity (U) of LipA (A) and

LipB (B) were measured at increased concentration of purified enzymes. Assay

reactions were conducted as described in chapter 2.

205

206

Fig. A.3.

The effect of commercially available detergents on the catalytic activity of LipA (A)

and LipB (B). Detergents were autoclaved prior to assays. Control is the enzyme assay

without detergent. 100%

207

208

Fig. 4.A.

SDS-PAGE of partially purified LipA and LipB. The Protein was first precipitated from

the supernatant by cold acetone, then separated and partially purified by Octyl

Sepharose chromatography. Gels were stained with GelCode® Blue Stain Reagent

(PIERCE).

209

210