1. Introduction

Introduction

2

O R1

O

OR2

O

O R3

O

OH

HO

OH

3H2O

Lipase

R1COOH

R2COOH

R3COOH

+

Triacylglycerol Glycerol Fatty acids

The biological relevance and large variability of lipids has led to the evolution

of a vast variety of lipid degrading enzymes throughout all kingdoms of life. Among

them, esterases, belonging to the group of enzymes that catalyze the cleavage of

chemical linkages by the addition of a water molecule are considered to be the most

important catalysts due to their widespread biological functions and biotechnological

potential (Bornscheuer, 2002b). Esterases (EC 3.1.1.1) have attracted considerable

attention because of their applications for new compound synthesis used in food,

pharmaceutical and chemical industries (Gupta et al., 2004). Esterases catalyze the

hydrolysis of ester bonds of lipids and other organic compounds, although they are

able to hydrolyze non-ester bonds (Sterner, 1999). Generally, esterases are specific for

either the alcohol or acid moiety and therefore, can be either alcohol or carboxylic

ester hydrolases (Fojan et al., 2000). Esterases have been classified by the

recommendations of Nomenclature Committee of the International Union of

Biochemistry and Molecular Biology (IUBMB) according to the specific bond,

moiety and substrate they hydrolyze. The carboxylic ester hydrolases, which act on

acylglycerols to liberate free fatty acids (FFA) and glycerol, are classified as lipases

(Gupta et al., 2004; Testa & Mayer, 2006).

Lipase

Lipases (acylglycerol acylhydrolases EC 3.1.1.3) are ubiquitous enzymes

widely distributed in the microbial, plant and animal kingdom (Gilbert, 1993; Jaeger

et al., 1994). They are of considerable physiological significance with industrial

potential. Lipases are primarily responsible for the hydrolysis of acylglycerides during

lipid processing and principally catalyze the hydrolysis of ester bonds in mono, di and

triacylglycerols (TAGs) (Figure 1. 1). They also possess the unique feature of

carrying out reactions such as esterification and transesterification involving water-

insoluble esters (Brockman et al., 1988).

Figure 1. 1. Hydrolysis of triacylglyceride by lipase.

Introduction

3

Occurrence of lipases

Lipase occurs widely in nature, throughout the Earth‟s flora and fauna.

However, they are found more abundantly in microbial flora comprising of bacteria,

fungi and yeast and most of them are extracellular (Macrae & Hammond, 1985;

Davranov, 1994; Jaeger et al., 1994). This ready availability has created an enormous

spin-off with respect to enantioselective hydrolysis and formation of carboxyl esters.

With the advent of genetic engineering techniques, an increasing number of lipases

are commercially produced from recombinant bacteria and yeasts. Although a number

of lipase producing bacterial sources are available, only a few are commercially

exploited as wild or recombinant strains (Jaeger et al., 1994; Palekar et al., 2000). Of

these, the important ones include the genera Achromobacter, Alcaligenes,

Arthrobacter, Bacillus, Burkholderia, Chromobacterium and Pseudomonas. The

lipases from Pseudomonas are widely used for a variety of biotechnological

applications (Jaeger et al., 1994; Pandey et al., 1999; Beisson et al., 2000). Fungal

lipases are extracellular enzymes, which facilitate the downstream processing (Jaeger

et al., 1999; Sharma et al., 2001). Some of the major lipase producing fungi are from

the genera Mucor, Rhizopus, Geotrichum, Rhizomucor, Aspergillus, Humicola,

Candida and Penicillium (Tan et al., 2003; Larios et al., 2004).

Lipases have been isolated from many insects, fish and mammals. These

lipases play an important role in digestion of lipids in biological system (Walton et al.,

1984). The most important source of animal lipase is the pancreas of cattle, sheep,

hogs and pigs (Vakhlu & Kour, 2006).

In plants, lipases are found in oleaginous seeds, especially in those which

contains large amount of TAGs. These include: corn (Zea mays), sunflower

(Helianthus annuus), rape seed (Brassica napus), castor bean (Ricinus communis) and

sesame (Sesamum indicum). In seeds, lipolytic activity normally appears during the

germination and found in sub-cellular compartments, either free or associated with

organelles. Most of the TAG lipases in plants are membrane bound associated with

either oil body, glyoxysome or microsomal fractions depending upon the species

(Huang et al., 1988; Mukherjee, 1994). Lipases from rape seed, castor bean, maize

and rice have been solubilized and extensively purified (Mukherjee, 1994). The

members of the Poaceae plant families are also a rich source of lipase. Cereal grains

contain up to 10 % lipids, normally located in the embryo and aleurone layer. In

wheat, the lipolytic activity (75-80 %) is located in the bran and 20-25 % in the germ

Introduction

4

(Tavener & Laidman, 1972). Other sources of plant lipases are laticifers, where

lipolytic activity is found in the insoluble fraction of latex. As early as 1935, lipolytic

activity in the latex in several plant families, including Asclepiadaceae, Moraceae,

Apocynaceae, Euphorbiaceae, Caricaceae and Bromeliacea was reported (Rivera et

al., 2012). These lipases are mostly trapped in the insoluble fraction, therefore

considered as naturally immobilized enzymes (Maria et al., 2006).

Reactions catalyzed by lipase

Lipase act on TAGs as well on a wide variety of natural and artificial

substrates and therefore catalyzes diversified reactions. The lipase catalyzed reactions

are classified into two main categories:

1. Hydrolysis:

RCOOR' + H2O RCOOH + R'OH

2. Synthesis: Reactions under this category can be further classified into:

a) Esterification

RCOOH + R1OH RCOOR

1 + H2O

b) Interesterification

RCOOR1 + R

2COOR

3 RCOOR

3 + R

2COOR

1

c) Alcoholysis

RCOOR1 + R

2OH RCOOR

2 + R

1OH

d) Acidolysis

RCOOR1 + R

2COOH R

2COOR

1 + RCOOH

e) Amonolysis

RCOOR1 + R

2NH2 RCOOR

2NH + R

1OH

Lipases catalyze the hydrolysis of fatty acyl esters or TAGs into FFA. In a

water-free environment, lipases catalyze a synthesis reaction between a carboxylic

acid and an alcohol to form an ester called esterification. In an interesterification

reaction, an acyl residue of one ester is transferred to another ester in the absence of

water. In a transesterification reaction, an acyl residue is either transferred to an

alcohol (alcoholysis) or to a carbonic acid (acidolysis). The product of an

ammonolysis reaction is an amide and an alcohol.

Introduction

5

CH2COOR

C

CH2COOR

RCOO H sn2

sn1

sn3

Specificity of lipases

Generally, lipases are classified into five different groups. 1) Substrate

specific, 2) regioselective (positional specific), 3) non-specific, 4) fatty acid specific

and 5) stereospecific lipases.

1. Substrate specific lipases

Substrate specificity is defined as the ability of a lipase to hydrolyze

preferentially a particular glycerol ester. These enzymes not only catalyze the

hydrolysis of TAGs but also hydrolyze di- and monoacylglycerols and even

phospholipids, in the case of phospholipases. TAGs are preferred substrate for

majority of plant and microbial lipases. Lipases show specificity not only with respect

to fatty acids but also with respect to alcohol part of the substrates (Jensen et al.,

1983). Lipase activities with different classes of alcohols are in the order: primary >

secondary > tertiary (Kuo & Parkin, 1993). Tertiary alcohols and their esters are poor

substrates for lipases (O'Hagan & Zaidi, 1994; Bosley et al., 1997).

2. Regioselective (positional specific) lipases

Regioselectivity is the ability of lipases to distinguish between the two

external positions (sn1 or sn3 position) and the internal position (sn2 position) of

TAG backbone (Figure 1. 2).

a) 1, 3 specific lipases catalyze reactions of the primary hydroxyl groups of TAG

(Saxena et al., 1999; Ribeiro et al., 2011). Thus, these preferentially release

fatty acids from positions sn1 and sn3 over the sn2 position to give an

equimolar mixture of 1, 2- and 2, 3-diacylglycerols. The subsequent

hydrolysis leads to 2-monoacylglycerol (Figure 1. 3).

b) True sn2 regioselectivity is very rare and Candida antarctica lipase A

(Rogalska et al., 1993) and a phospholipase A2 from rice bran (Bhardwaj et

al., 2001) has been reported that preferentially hydrolyzes the sn2 position of

the TAG backbone.

Figure 1. 2. Representation of sn-nomenclature for TAGs.

Introduction

6

CH2COOR

C

CH2COOR

RCOO H

H2O

CH2OH

C

CH2COOR

RCOO H + RCOOH

H2O

CH2OH

C

CH2OH

RCOO H + RCOOH

Triacylglycerol 1,2 (2,3)-Diacylglycerol 2-Monoacylglycerol

CH2COOR

C

CH2COOR

RCOO H

3H2O

CH2OH

C

CH2OH

HO + 3RCOOH

Triacylglycerol Glycerol Fatty acids

Non-specific lipases

1,3-specific lipases

A

B

Figure 1. 3. Reactions catalyzed by A) 1, 3 specific lipases and B) non-specific lipases.

3. Non-specific lipases

These enzymes catalyze the complete hydrolysis of TAGs and thus can

remove fatty acid from any position of the TAG (Macrae & Hammond, 1985; Saxena

et al., 1999) resulting in complete breakdown to glycerol and FFA (Figure 1. 3).

Diacylglycerides and monoacylglycerides are formed as intermediates in the reaction

mixture (Saxena et al., 1999). These intermediates are hydrolyzed more rapidly than

TAG and do not accumulate in the reaction.

4. Fatty acid specific lipases

These are either specific for a particular type of fatty acid or, more frequently,

for a specific group of fatty acids. Such lipases hydrolyze TAG esters of these acids

regardless of their position on the glycerol backbone. They hydrolyze fatty acid esters

located at any position on the TAG. Some lipases prefer hydrolysis of those esters,

which are formed from long-chain fatty acids with double bonds (Jensen, 1974).

Aspergillus flavus lipase shows a greater preference for tricaprin compared to

triolein (Long et al., 1998). Candida rugosa and Rhizomucor miehei lipase have

strong preference for oleic over elaidic acid whereas C. antarctica lipase A prefers

elaidic over oleic acid (Borgdorf & Warwel, 1999). Oat seed lipase discriminates

against tripetroselinin as compared to triolein, trilinolein and tri--linolein (Piazza et

al., 1992). Lipases also exhibit fatty acid chain length specificity. Most of the lipases

prefer esters of medium (C4) to long-chain (C16) saturated fatty acids with a few

exceptions. Lipases not only accommodate TAGs and aliphatic esters but also

compounds like alicyclic, bicyclic and aromatic esters and even esters based on

organometallic sandwich compounds (Chen & Sih, 1989; Schmid & Verger, 1998).

Introduction

7

5. Stereospecific lipases

Stereospecificity is defined as the ability of a lipase to distinguish between sn1

and sn3 position of the TAG. Lipases may show specific or insignificant

stereoselectivity. The stereoselectivity of the same enzyme may vary according to the

structure of the substrate (Sonnet, 1988). A change in stereoselectivity of lipases can

occur with different substrates e.g. stereoselectivity for sn3 position for trioctanoin

changes to sn1 position for triolein (Rogalska et al., 1993).

Along with the above described specificities, lipases are also known to display

enantioselectivity and condition promiscuity.

Enantioselectivity

Enantioselectivity is the ability to discriminate enantiomers in a racemic

mixture. The enantio specificities of lipases vary according to the substrate and this

variation is connected to the chemical nature of the ester. Enantioselective processes

based on lipase catalysis include the synthesis of chiral amines, catalysed by the lipase

from Burkholderia plantarii (Balkenhohl et al., 1997) and Serratia marcescens lipase-

based production of (2R, 3S)-3-(4-methoxyphenyl) methyl glycidate, which is used in

the synthesis of the calcium antagonist DiltiazemTM

(Shibatani et al., 1990).

Condition promiscuity

Organic solvents have been found to influence selectivity of lipase catalyzed

reactions (Rubio et al., 1991; Hirose et al., 1992). Solvents can affect regioselectivity

and prochiral selectivity. Prochiral selectivity is the ratio of the rate of accumulation

of major enantiomer over that of minor enantiomer. In organic solvent, lipases carry

out the reverse of hydrolysis, i.e., synthesis and catalyze esterification,

transesterification and interesterification (Klibanov, 1990; Rodrigues & Fernandez-

Lafuente, 2010). Water-organic solvent two-phase systems have been employed

successfully to transform highly lipophilic substrates such as steroids, fats and

alkenes. The esterification and interesterification activities of Rhizopus nivea lipase

and R. miehei lipase was studied in three different types of organic media. Biphasic

systems gave better results with esterification reactions as compared to

interesterification reactions (Antonini et al., 1981; Kim et al., 1984; Brink & Tramper,

1985).

Introduction

8

Purification strategies for lipase

Various purification strategies used for lipases have been reviewed, clearly

highlighting the importance of designing optimal purification schemes (Aires-Barros

& Cabral, 1991; Taipa et al., 1992; Palekar et al., 2000; Sharma et al., 2001; Saxena et

al., 2003). Purification procedures are usually a multistep series of non-specific

techniques such as ammonium sulphate precipitation, gel filtration and ion exchange

chromatography. Affinity chromatographic procedures have been increasingly used,

facilitating the purification of lipases.

Most of the microbial lipases are extracellular, being secreted into the culture

medium. Purification of these enzymes involves removal of cells from the culture

broth, either by centrifugation or filtration. The cell-free supernatant obtained is then

concentrated either by ultrafiltration or extraction with organic solvents or ammonium

sulphate precipitation. About 80 % of the purification schemes apply a precipitation

step, 60 % of which use ammonium sulphate and 35 % use ethanol, acetone or an

acid. These are followed by a combination of several chromatographic methods, such

as gel filtration and affinity chromatography. Strong ion exchangers based on

triethylaminoethyl groups (Veeraragavan et al., 1990) and Q-sepharose (Menge et al.,

1990) are becoming popular in lipase purification. Concavalin A (Con A) and heparin

were employed to purify fungal and mammalian lipases based on their glycoprotein

nature (Tombs & Blake, 1982; Aires-Barros & Cabral, 1991). The final step of gel

filtration normally yields a homogeneous product (Scopes, 1994; Vasudevan, 2004).

Novel purification techniques like membrane processing, immuno-purification,

hydrophobic interaction chromatography and aqueous two phase systems have also

been employed (Saxena et al., 2003).

Since lipases are known to be hydrophobic in nature with large hydrophobic

surfaces around the active site, the purification of lipases is achieved by opting for

hydrophobic interaction chromatography. The most popular hydrophobic adsorbents

being used are butyl, octyl or phenyl functional groups. The application of

hydrophobic interaction chromatography in combination with other chromatographic

procedures in purification of lipase from various sources has been extensively

reviewed (Taipa et al., 1992). A hydrophobic lipase from Pseudomonas sp. was

purified with high yield of 56 % and purity of 159 fold, using hydrophobic interaction

chromatography on octyl Sepharose in combination with Q-Sepharose (Kordel et al.,

1991).

Introduction

9

The hydrophobic nature of lipases is exploited in the aqueous two-phase

bioseparation by using detergents or surfactants during the purification. A detergent

based aqueous two-phase system used to purify Pseudomonas cepacia lipase

indicated that all prokaryotic lipases show a preference for the detergent based

coacervate phase (Terstappen et al., 1992). The selective separation and purification

of a lipase B from Chromobacterium viscosum was achieved in AOT-isooctane

micellar solution (Vicente et al., 1990).

Lipase assay methods

An efficient assay procedure is of paramount interest for screening and

identifying lipase producing species/clones from a diverse population. Lipases can be

viewed both as lipolytic and esterolytic enzymes, and can thus be assayed by

monitoring the release of either fatty acids or glycerol from TAGs or fatty acid esters.

Gel diffusion assay has been used to screen lipases in culture supernatants.

Lipase activity is detected by the appearance of a clear zone in the gel supplemented

with mechanical emulsions of TAG (Lawrence et al., 1967; Stead, 1986).

Alternatively, the released fatty acids is detected by addition of pH indicators like

Victoria Blue and Nile Blue Sulphate, which form complexes with the acids and a

coloured zone is produced. Addition of a fluorophore such as Rhodamine B,

complexes with FFA producing an orange-pink fluorescence under UV irradiation

(Kouker & Jaeger, 1987).

Titrimetry is among the oldest and most widely used quantitative methods for

lipase assay. This is a reliable method for characterizing lipase action, specificity and

measuring kinetic parameters (Ferrato et al., 1997). The release of FFA using a

mechanically stirred emulsion of natural or synthetic TAGs is measured either by

manual titration with NaOH to a phenolphthalein end point or by pH-stat titration to

constant end point value (Beisson et al., 2000). The pH-stat method is a highly

sensitive and quantitative method that can measure the release of even 1 µmol of

released fatty acid/min (Jaeger et al., 1994).

Spectrophotometric assay methods are widely used as they are fast and simple.

p-nitrophenyl esters of various chain length fatty acids are commonly used as

substrates and the release of p-nitrophenol is measured spectrophotometrically at 410

nm (Jaeger et al., 1999). However, care is required when interpreting activity

measurements in crude samples using certain short chain p-nitrophenyl esters as

Introduction

10

substrates, as they can be hydrolyzed by other enzymes or basic solutions (Beisson et

al., 2000). Colorimetric assays are based on hydrolysis of the colourless ester -

naphthyl esters to yield coloured -naphthol, which is quantifiable at 560 nm

(Nachlas & Blackburn, 1958; Degrassi et al., 1999; Gandolfi et al., 2000).

Alternatively, TAG derivative 1,2-O-dilauryl-rac-glycero-3-glutaric acid resorufin

ester can be used, which on hydrolysis yields resorufin that is determined at 572 nm

(Jaeger et al., 1999).

Fluorescence assays involve measurement of the fluorescent fatty acid

released because of lipase activity. They are usually based on the use of TAG

analogues marked with fluorophores such as dansyl, resorufin or 4-

methylumbelliferone groups. Lipolysis of these analogues releases the fluorophores,

which can be detected fluorimetrically (Beisson et al., 2000; Prim et al., 2000). Other

approaches include the detection of fatty acids released from TAGs by fluorimetric

determination of Rhodamine-B fatty acid complexes (Beisson et al., 2000).

A quantitative analysis of the released FFA from TAGs can be carried out by

thin layer chromatography using autoradiographic methods with radiolabelled TAGs.

In GC, the released fatty acids are converted into their methyl esters and subjected to

quantification.

The oil-drop tensiometer presents the unique advantage of being able to

monitor lipase activities on natural long-chain TAGs in a closely controlled oil/water

interface. Lipase catalyzed hydrolysis of the lipid monolayer results in changes on the

surface pressure, which is readjusted automatically by a computer controlled Barostat

(Jaeger et al., 1999). Conductimetric, turbidimetric and IR assays are also used to

detect hydrolysis or synthesis catalyzed by lipases (Gupta et al., 2003).

Distinguishing a lipase from an esterase

Lipase and lipolytic enzymes are generic terms that include two main groups

of enzymes: carboxylesterases and “true” lipases, which differ with respect to their

preference for short or long chain substrates, respectively. However, enzymes such as

cutinases and phospholipases which can hydrolyze acylglycerols are also considered

as lipolytic enzymes by some authors (Longhi & Cambillau, 1999; Fojan et al., 2000).

Several criteria have been employed to distinguish TAG lipases or true lipases (EC

3.1.1.3) from esterases (EC 3.1.1.1). However, substrate specificity and gene

sequence discrimination are novel criteria for this distinction. Several of the known

Introduction

11

lipase gene sequences are homologous and have led to the classification of different

families of lipases/esterases (Arpagaus et al., 1990; Derewenda & Derewenda, 1991).

Despite the molecular structure and catalytic mechanism of lipase and esterase being

similar, a lipase is distinguished from an esterase by its preference for substrates and

interfacial activation and/or presence of a lid (Fojan et al., 2000; Bornscheuer, 2002b;

Ewis et al., 2004).

Microbial lipases

Microbial lipases today occupy a place of prominence among biocatalysts

owing to their ability to catalyze a wide variety of reactions in aqueous and non-

aqueous media. These lipases have gained special and industrial attention due to their

exquisite chemoselectivity, regioselectivity and stereoselectivity (Dutra et al., 2008;

Griebeler et al., 2011). Microorganisms including bacteria, yeast and fungi are

potential producers of extracellular lipases (Abada, 2008). Microbial lipases are

readily available in large quantities as they can be produced in high yields (Pandey et

al., 1999). The purification of lipases from various sources has enabled successful

sequence determination. The crystal structures of many lipases have been solved

facilitating the understanding of their unique structure-function relationships and

design of rational engineering strategies (Aravindan et al., 2007). Lipase producing

microorganisms have been found in different habitats such as industrial wastes,

vegetable oil processing factories, dairy plants and soil contaminated with oil and

oilseeds (Sharma et al., 2001). Among the bacterial lipases, the Bacillus exhibit

interesting properties and make them potential candidates for biotechnological

applications. In addition, Pseudomonas sp., Pseudomonas aeruginosa, Burkholderia

multivorans, Burkholderia cepacia, and Staphylococcus caseolyticus are also reported

as bacterial lipase producers (Treichel et al., 2010).

Most fungal lipases are of considerable commercial importance for bulk

production. Although a number of lipase producing fungi are recognized, the most

important belong to the genera Rhizopus sp., Aspergillus sp., Penicillium sp.,

Geotrichum sp., Mucor sp., and Rhizomucor sp. (Singh & Mukhopadhyay, 2012). C.

rugosa lipases and their isoforms have great significance for their diverse

biotechnological potential (Pandey et al., 1999). The main terrestrial species of yeasts

Introduction

12

that were found to produce lipase are Candida sp., Yarrowia lipolytica, Rhodotorula

glutinis, Rhodotorula pilimanae, Pichia bispora, Pichia mexicana, Pichia sivicola,

Pichia xylosa, Saccharomycopsis crataegenesis and Trichosporon asteroids. The

genes encoding lipase in Candida sp., Geotrichum sp., Trichosporon sp., and Y.

lipolytica have been cloned and over expressed (Wang et al., 2007). The Y. lipolytica

lipase, YLLIP2 was shown to be an ideal candidate for enzyme replacement therapy

due to its unique biochemical properties. It shows highest activity at low pH values

and is not repressed by bile salts. YLLIP2 belong to the same gene family as

Thermomyces lanuginosus lipase, a well known lipase with many applications in the

field of detergents and biotechnological applications (Aloulou et al., 2007).

Mammalian lipases

Mammalian lipases can be distinguished into three groups: the lipases secreted

into the digestive tract by specialized organs, the tissue lipases and the milk lipases

(Desnuelle, 1972). Classification of the digestive enzymes was proposed based on the

tissular and cellular origin, site of lipolytic action and substrate specificity (Table 1. 1)

(Gargouri, 1989). Isolation and purification techniques used in the recovery of lipases

from mammalian sources have been reviewed (Taipa et al., 1992). Purification of

mammalian lipases allowed the determination of the primary amino acid sequence of

several important lipases such as mammalian pancreatic and gastric lipases (Verger,

1984; Gargouri, 1989). The major energy reserve in animals is stored in the form of

lipids constituted in adipocytes. Mobilization of TAGs stored in adipocytes is tightly

regulated by hormones and requires activation of lipolytic enzymes. Upon

stimulation, protein kinase A phosphorylate hormone sensitive lipase, which is then

translocated to lipid particles where they catalyze release of fatty acids from TAG.

This enzyme not only hydrolyzed TAG but exhibited even higher hydrolytic activity

for diacylglycerol and cholesterol oleate (Osterlund et al., 1996; Ali et al., 2005). The

purification of pancreatic lipases from rat, bovine, sheep and horse has been

extensively reported (Verger, 1984). The physiological role of liver lipases that

degrade TAG in cytosolic lipid droplets, endoplasmic reticulum, lysosomes and along

the secretory route has been reviewed (Quiroga & Lehner, 2012).

Introduction

13

Table 1. 1. Classification of digestive lipases.

Site of action

Stomach Small intestine

Generic

names Acid lipases

Colipase

dependent lipases

Bile salt dependent lipases

Tissular

origin

Tongue

Pharynx

Stomach

Exocrine

pancreas

Human milk

Exocrine pancreas

Specific

names

Lingual lipase

Salivary lipase

Pharyngeal lipase

Gastric lipase

Preduodenal lipase

Pancreatic lipase

Bile salt-stimulated lipase

Carboxyl-ester lipase

Non-specific lipase

Plant lipases

The knowledge of plant lipases is limited compared to those of animal and

microbial origin. Plant lipases can be broadly classified into: 1). TAG lipases or

“true” lipases (EC 3.1.1.3) that hydrolyze TAGs, 2) non-specific lipid acylhydrolases

exhibiting combined action of various lipases, such as phospholipase A1 (EC

3.1.1.32), A2 (EC 3.1.1.4), B (EC 3.1.1.5), glycolipase, sulpholipase and

monoacylglycerol lipase occurring in diverse plant tissues and 3) phospholipase C

(EC 3.1.4.3) and D (EC 3.1.4.4), the latter being more widely distributed in plants

(Borgston & Brockman, 1984). These lipases represent an important group of

hydrolases that combine competitive prices with a wide versatility and stability in

organic media (Caro et al., 2002). Lipases from plant families like Euphorbiaceae

(Giordani et al., 1991; Moulin et al., 1994; Palocci et al., 2003; Villeneuve et al.,

2005), Asclepiadaceae, (Giordani et al., 1991), Brassicaceae (Hills et al., 1990) and

Caricaceae (Giordani et al., 1991; Dhuique-Mayer et al., 2001; Maria et al., 2006)

have been described as useful biocatalysts for several applications.

Oilseed lipases are usually more active with endogenous TAGs containing

fatty acids of varying chain length. Certain seed lipases show selectivity for the

dominant fatty acids in the seeds (Hellyer et al., 1999). For e.g., palm tree lipase is

specific for tricaproin or trilaurin; Vernonia sp. lipase for trivernolein; castor bean

lipase for tricaproin or trilaurin and elm lipase for tricaproin. Corn lipase presented

greater activity with the TAGs containing oleic and linolenic acids, which are the

main constituents of corn oil (Lin & Huang, 1984). Depending on the plant species,

Introduction

14

the lipase may be located in the membrane of the lipid bodies or in other cellular

compartments (Borgston & Brockman, 1984). The lipases of the following oil seeds

have been most widely studied with respect to extraction and characterization: beans

(Enujiugha et al., 2004), sunflower seeds (Sadeghipour & Bhatla, 2003; Sagiroglu &

Arabaci, 2005); linseed (Sammaour, 2005); peanuts (Huang & Moreau, 1978) and

cotton seeds (Rakhimov et al., 1970), although lipases from other oilseed sources are

currently being investigated.

Lipases from various beans have been well characterized. A Ca2+

dependent

thermostable lipase from Africa bean seeds (Pentaclethra macrophylla) showed

greater activity with oils containing short chain fatty acids (Enujiugha et al., 2004). A

study on French bean lipase showed that the enzyme presented greater activity at pH

7.0 and that the addition of Ca2+

had an inhibitory effect, whereas the addition of the

emulsifier Tween-20 resulted in a four-fold increase in enzyme activity. The

specificity of French bean lipase against various substrate showed that triacetin was

the best substrate (Kermasha & Van de Voort, 1986). Castor beans contain a lipase

with some peculiar characteristics. The enzyme has an optimum pH of 4.5 and was

inactivated at pH values above 6.0 at 30 C. The specificity was observed towards

medium chain fatty acids and also for unsaturated fatty acids. In addition, it showed

some regioselectivity for fatty acids at the positions sn1 and sn2 (Eastmond, 2004).

The degradation of stored lipids, in correlation with the seed proteins was

studied during germination of sunflower (Helianthus annuus) seeds. Lipolytic activity

increased in the seeds grown in sunlight (Sadeghipour & Bhatla, 2003). A 22,000 Da

sunflower seed lipase showed preference for TAGs with mono-unsaturated fatty acids

(Sagiroglu & Arabaci, 2005). A lipase form germinated canola (Brassica napus) seeds

was activated in the presence of Ca2+

and Bi3+

ions by 165 % and 124 % respectively

(Sana et al., 2004). The enzyme showed high activity with trierucic, tripalmitate and

4-methyl-umbeliferyl oleate as substrates (Lin et al., 1986). Among the grain lipases

studied (wheat, linseed, barley and canola), the canola seed lipase showed highest

degree of flavor formation producing (Z)-3-hexen-1-yl butyrate and (Z)-3-hexen-1-yl

caproate with an efficiency of about 96 %.

A partially purified lipase from Barbados nuts (Jatropha curcas) seeds showed

a high activity with triolein at a pH of 7.4. Addition of Ca2+

increased the enzyme

activity by 130 % whereas Fe2+

inhibited lipase activity (Abigor et al., 2002).

Although two esterases and a lipase were identified in Barbados nuts, only the lipase

was observed during germination period (Staubmann et al., 1999).

Introduction

15

The lipase activity of lupin (Lupinus luteus) cultivated in vitro showed higher

activity when sugar was absent in the medium (Borek et al., 2006). Lupin seed lipase

showed a greater specificity for fatty acids at positions sn1 and sn2 of the TAGs of

lupin seed oil and was more active with saturated than unsaturated fatty acids (Sanz &

Olias, 1990). A partially purified lipase from almond (Amygdalus communis) seed

showed a wide range of specificity towards natural oils (Yesiloglu & Baskurt, 2008).

Wheat (Triticum aestivum) germ lipase originally described by Singer and

Hofstee (1948) is commercially available. Some authors have classified wheat lipase

as an esterase, however studies carried out using triolein as the substrate has shown

good activity rendering it a lipase (Jing et al., 2003; Kapranchikov et al., 2004;

Korneeva et al., 2008). Lipase activity was high in bran compared to the whole kernel

and highly related to the pool of FFA in the stored wheat (Rose & Pike, 2006).

Purified wheat lipase, a 143,000 ± 2,000 Da protein was thermostable (Kapranchikov

et al., 2004). The role of Ser-OH group in the catalytic action of wheat germ lipase

has been demonstrated (Korneeva et al., 2008).

Corn (Zea mays) grains lipase was induced only two days after germination

and decreased with depletion of the stored lipid. The corn seed lipase showed higher

activity with TAGs that contained linoleic and oleic acids (Lin et al., 1984; Huang et

al., 1988). The synthesis of esters by seed lipases, precipitated with ammonium sulfate

was studied in a medium containing organic solvents (Liaquat & Apenten, 2000).

Corn lipase showed better activity with short chain fatty acids in the following order:

acetic > butyric > caproic acids in an organic medium using isopentanol. A lipase

from dog was cloned and expressed in transgenic corn seed (Zhong et al., 2007). The

stability studies in different surfactants showed that the recombinant enzyme was

stable.

The lipase of oats (Avena fatua) is localized on the surface of oat caryopsis

(Martin & Peers, 1953) and exists as four isoenzymes that are heat stable. The

resistance to high temperatures and activity at alkaline pH values promotes oat lipases

for possible industrial applications (Mohamed et al., 2000). A lipase identified in

germinated barley (Hordeum vulgare) grains, showed maximum activity two days

after germination (Kubicka et al., 2000). The seeds of sorghum (Sorghum bicolor)

exhibit lipase activity during grain malting and mashing, which decreased during

steeping. However, there was a several fold increase of lipase activity during the

course of germination (Nwanguma et al., 1996). The most widely studied seed lipases

and their biochemical properties are summarized (Table 1. 2).

Introduction

16

Table 1. 2. The most studied seed lipases and their biochemical properties.

Lipase

source

Optimum

pH

Optimum

temperature

(C)

Specificity Activators Inhibitors

Africa

bean 7.0 30 - Ca

2+ EDTA

French

bean 7.0 35 - Tween-20 Ca

2+

Castor

bean 4.5 30 sn1, sn2 Ca

2+

Chloromercuri

-benzoic acid

Rapeseed 7.0 37 - Bi2+

, Ca2+

Fe3+

, Fe2+

,

Zn2+

, Hg2+

and

Cu2+

Barbados

nut 7.5 45 - Ca

2+, Mg

2+ Fe

2+

Lupin 5.0 40 sn1, sn2 Ca2+

, Mg2+

, K+ -

French

peanut 8.0 65 - Ca

2+, Mg

2+

Hg2+

, Mn2+

,

Zn2+

and Al3+

Almond 8.5 65 -

Ca2+

, Fe2+

,

Mn2+

, Co2+

and Ba2+

Mg2+

, Cu2+

and Ni2+

Laurel 8.0 50 -

Ca2+

, Mg2+

,

Co2+

, Cu2+

and

Fe2+

-

Black-

Cumin 6.0 45 - - -

Rice

11.0

7.5

7.5

80

37

27

sn2

sn1, sn3

sn1, sn3

- -

Wheat 8.0 37 - - -

Oat 9.0 65-75 - Ba2+

, Ca2+

Mn2+

and Zn2+

Coconut 8.5 30-40 sn1, sn2 - -

Introduction

17

Physiological role of lipases

The physiological functions of lipases are not yet clear for many of them,

although they seem to be involved in the bioconversion of lipids (Pandey et al., 1999).

Microbial lipases display broad substrate specificity, a property that seems to have

evolved to ensure the access of lipase producing microorganisms to diverse carbon

sources during plant cell wall degradation or during the recycling of lipid-containing

nutrients (Gunstone, 1999; Sharma et al., 2001). Some lipases are involved in the

turnover of membrane lipids, adapting to environmental changes by altering the cell

membrane composition. They are also involved in cell signaling in controlled

destruction of intracellular vacuoles, in cytolysis (Schmid & Verger, 1998). In

eukaryotes, lipases are involved in various stages of lipid metabolism including fat

digestion, absorption, reconstitution and lipoprotein metabolism (Desnuelle et al.,

1986).

Physiological role of lipases in plants are mainly involved in the regulation of

plant development, morphogenesis, synthesis of secondary metabolites and defense

response (Ling et al., 2006). TAG hydrolysis in plants play a pivotal role providing

the carbon skeletons and energy for post-germination growth. Lipases catalyze the

initial step in TAG breakdown at oil/water interface to yield FFA and glycerol

(Quettier & Eastmond, 2009). In most plants the lipase activities are detectable upon

germination and increase concomitantly with the disappearance of TAGs. Depending

on the species, these lipases are often membrane associated and can be found in oil

body, glyoxysome or microsomal fractions of seed extracts (Huang, 1992; Mukherjee,

1994). Positional cloning of a sugar-dependent1 (sdp1) that encodes a lipase, has

revealed the molecular identity of an oil body associated TAG lipase that is

responsible for catalyzing the initial step in storage oil mobilization in Arabidopsis

seeds (Eastmond, 2006). Genetic evidence has shown that TAG lipases are required

for oil breakdown (Athenstaedt & Daum, 2003; Zimmermann et al., 2004; Gronke et

al., 2005). An Arabidopsis GDSL lipase GLIP1 possessed lipase and anti-microbial

activities that directly disrupted fungal spore integrity, and in association with

ethylene signaling played a key role in plant resistance to Alternaria brassicicola

(Ling et al., 2006). In addition to post-germinative TAG breakdown, some

degradation has been observed during late seed development in several plant species.

The TAG content in oilseed rape (Brassica napus) embryos decreased by ~10 % by

the time desiccation was complete. The precise physiological role of this TAG

breakdown is not known (Chia et al., 2005).

Introduction

18

Three-dimensional structure of lipase

The three-dimensional structure of many lipases has been determined by X-ray

crystallography and nuclear magnetic resonance (NMR) spectroscopy (Breg et al.,

1995; Fojan et al., 2000; Chen et al., 2009; Angkawidjaja et al., 2010). The lipases

whose X-ray structures have been determined are listed in Table 1. 3. The following

structural features are common to lipases:

1. Lipases are members of “/ hydrolase fold” family. Although different lipases

may display low sequence similarity, they show structural similarity which

comprises of a core, predominantly parallel strands surrounded by helices

(Van et al., 2001; Carr & Ollis, 2009; Widmann et al., 2010). The canonical /

hydrolase fold consists of a central parallel sheet of eight strands with the

second strand antiparallel. The parallel strands 3 to 8 are connected by

helices, which pack on either side of the central sheet (Figure 1. 4).

Table 1. 3. Lipases whose X-ray structures have been determined.

Lipase source PDB ID

Bacterial

Bacillus thermocatenulatus 2W22

Burkholderia glumae 1TAH

Burkholderia cepacia 1OIL

Bacillus subtilis 2QXU

Chromobacterium viscosum 1CVL

Fungal

Candida rugosa 1GZ7

Candida antarctica A

3GUU

Candida antarctica B 1TCC

Geotrichum candidum 1THG

Penicillium camembertii 1TIA

Penicillium expansum 3G7N

Rhizomucor miehei 4TGL

Rhizopus oryzae 1TIC

Rhizopus niveus 1LGY

Thermomyces lanuginosus 1DT5

Yarrowia lipolytica 3O0D

Animal

Canis lupus familiaris 1RP1

Equus caballus 1HPL

Human pancreatic lipase 2OXE

Sus scrofa 1ETH

Introduction

19

Figure 1. 4. / hydrolase fold of lipases. The eight -sheets (1-8) are drawn as blue

arrows and the red cylinders represent the -helices (A-F). The distribution of the amino acids

(Ser, Asp, His) belonging to the catalytic triad are shown (Ollis et al., 1992). This figure is

reproduced with the kind permission of Dr. David L. Ollis.

2. The active site nucleophilic serine residue rests at a hairpin turn between a

strand and an helix in a highly conserved pentapeptide sequence GXSXG,

forming a characteristic -turn- motif named the „nucleophilic elbow‟ (Ollis et

al., 1992; Schrag & Cygler, 1997; Carrasco-Lopez et al., 2009).

3. The active site is formed by a catalytic triad consisting of amino acids Ser, His

and Asp/Glu (Brady et al., 1990; Winkler et al., 1990; Carrasco-Lopez et al.,

2009).

4. Presence of a lid or flap comprising of an amphiphilic helix peptide sequence

that covers the active site (Schmid & Verger, 1998).

The active site of lipase contain structures such as a fatty acid binding pocket

that facilitates the catalytic process, which is variable and responsible for

accommodation of acyl chain of the ester linkage that is to be hydrolyzed. An

additional binding pocket for the acyl chains of substrates like TAGs, which

contributes to anchoring of the substrate to active site of the enzyme during catalysis,

is reported (Jaeger et al., 1999). The lid covers the active site of the enzyme in the

absence of an interface. The lid may consist of a single helix, two helices or a loop

region, which is mainly hydrophobic in the region directed towards the active site and

hydrophilic on its external surface. The lid displays a variable position depending on

the physicochemical environment of the enzyme (Miled et al., 2003; Ericsson et al.,

2008). In the Bacillus thermocatenulatus lipase, the lid has a complex structure

involving a large percentage of amino acids of the protein and forms a double lid

(Carrasco-Lopez et al., 2009). Lipase B from C. antarctica has a very small and

Introduction

20

simple lid, which does not fully isolate the active center of the enzyme in the closed

form (Uppenberg et al., 1994). Guinea-pig lipase has a „mini-lid‟, which is composed

of only five amino acids (Hjorth et al., 1993). Structural rearrangement occur in the

presence of an interface displacing the lid, which is stabilized by hydrophilic

interactions with the adjacent enzyme surface allowing free access to the substrate and

solvent to the active site (Secundo et al., 2006; Gao et al., 2011).

Pleiss et al., (1998) subdivided lipases into three subgroups on the basis of the

geometry of the binding site i) lipases with a hydrophobic crevice like binding site

located near the protein surface (lipases from Rhizomucor and Rhizopus); ii) lipases

with a funnel-like binding site (lipases from C. antarctica, Pseudomonas and

mammalian pancreas) and iii) lipases with a tunnel-like binding site (lipase from C.

rugosa).

The basic features of the three-dimensional structure of the Rhizomucor miehei

lipase (RML) are described in detail (Brady et al., 1990; Derewenda et al., 1992) as

this is the lipase most often used for comparison. RML belongs to the / hydrolase

family. RML, a single polypeptide chain of 269 residues is folded into an unusual

singly wound -sheet domain and helical segments (Figure 1. 5). Three disulfide

bonds (residues 29-268, 40-43 and 235-244) stabilize the molecule. The catalytic triad

of RML is formed by Ser144

, Asp203

and His257

, similar in structure and function to the

analogous triad found in families of Ser proteases. The catalytic site is concealed

under a short amphipathic helix (residues 85 to 91), which acts as “lid”, opening the

active site when the enzyme is adsorbed at the oil-water interface. In the native

enzyme the “lid” is held in place by hydrophobic interactions. Three Ser residues

(Ser82-84

) and four other residues (92 to 95) placed at the ends of the helix form a

hinge, so that the helical part (Leu85

-Asp91

) can move as a rigid body in the course of

the activation process. A water molecule forms hydrogen bond with Ser144

of the

catalytic triad and Ser82

in the region of the “lid” helix and Leu145

(Derewenda et al.,

1990). These stabilize the correct conformation of the catalytic site in the closed and

open conformation, during the hydrolysis reaction of the ester bond. Ser82

plays a role

in substrate binding and is fixed by the water molecule in the correct orientation to

effectively bind the intruding substrate (Vasel et al., 1993). From the X-ray structure

of cocrystals of lipases and substrate analogues, there is strong evidence that the RML

undergoes a conformational rearrangement which renders the active site accessible to

the substrate (Brzozowski et al., 1991).

Introduction

21

A B

Free enzyme

NHN

HisHO

O-

Glu/Asp

Ser

O

O

O R

R'

NHN

HisO

OHGlu/Asp

Ser

O H

N

Gly/Ala

H

N

Gly/Ala

O

O R

R'

OO

R

R'

NHN

HisO

OHGlu/Asp

Ser

OH

N

Gly/Ala

H

N

Gly/Ala

OHO

RNH

N

HisO

OHGlu/Asp

Ser

OH

N

Gly/Ala

H

N

Gly/Ala

O

RN

N

HisO

OHGlu/Asp

Ser

OH

N

Gly/Ala

H

N

Gly/Ala

Acyl-enzyme complex R‘OH

O

HO R

Figure 1. 5. Structure of Rhizomucor miehei lipase in closed (A) and open (B)

conformation represented by space-filling model. The model is colored by decreasing

polarity. Upon opening of the lid, the catalytic triad (yellow) becomes accessible (B), and the

region binding to the interphase becomes significantly more polar (Schmid & Verger, 1998).

This figure is reproduced with the kind permission from John Wiley and Sons.

Figure 1. 6. Illustration of mechanism of substrate hydrolysis catalyzed by lipases.

Introduction

22

Mechanism of lipase action

The similarity of the catalytic triad found in lipases and proteases renders the

mechanism of lipase catalysis similar to that of a Ser protease, which involves

formation of two tetrahedral intermediates. Evidence for the mechanism of lipase

action has particularly come from crystallographic analysis of inhibitor-lipase

complexes. The mechanism of lipase action is represented in Figure 1. 6.

The mechanism involves the nucleophilic attack of the active site Ser-OH on

carbonyl carbon of the scissile ester bond of the substrate. The nucleophilic Ser

residue is activated by a proton transfer from the Ser-OH group to a neighboring

active His. Proton transfer is facilitated by the presence of the catalytic acid, which

precisely orients the imidazole ring of the His and partly neutralizes the charge.

Activation is followed by the attack of O- of the Ser hydroxyl group on the activated

carbonyl carbon of the susceptible lipid ester bond.

A transient tetrahedral intermediate is formed characterized by formation of

negative charge on the carbonyl oxygen atom of the scissile ester bond and the four

atoms bonded to the carbonyl carbon atom arranged as a tetrahedron. The

intermediate is stabilized by hydrogen bonds between the negatively charged carbonyl

oxygen (oxyanion) and at least two main chains -NH groups (oxyanion hole).

Subsequently, the additional proton of His is donated to the oxygen of the susceptible

ester bond, which is thus cleaved. At this stage, the acid component of the substrate is

esterified to the nucleophilic serine (the covalent intermediate), whereas the alcohol

component diffuses away.

In the deacylation step, the incoming water molecule hydrolyses the covalent

intermediate (acyl enzyme) and the acid component of the substrate is esterified to the

enzyme‟s Ser residue. The catalytic His activates the water molecule by withdrawing

a proton from it. The resulting OH- ion performs a nucleophilic attack on the carbonyl

carbon atom of the acyl group covalently attached to the Ser. Again, a transient

negatively charged tetrahedral intermediate is formed, which is stabilized by

interactions with the oxyanion hole.

The His residue donates its additional proton to the oxygen atom of the active

Ser residue, which breaks the ester bond between Ser and the acyl component,

releasing the acyl product. After diffusion of the acyl product, the enzyme is ready for

another round of catalysis.

Introduction

23

Interfacial activation

Lipases have evolved to deal with the biophysical properties of the interfacial

microenvironment where their substrates are to be found. In the past, the interfacial

influence has been described through the “quality of the interface” which determines

the behaviour of the lipase and outcome of the enzymatic reactions (Furuhashi et al.,

2007). The activity of lipase is low on monomeric substrates however, increases

dramatically as an aggregate supersubstrate-emulsion or a micellar solution is formed.

This phenomenon is called interfacial activation (Schmid & Verger, 1998). The

increase in enzymatic activity is triggered by structural rearrangements of the lipase

active site region as revealed from the crystal structures of lipase complexed with

small transition-state analogues (Jaeger et al., 1999; Overbeeke et al., 2000;

Angkawidjaja et al., 2010). The lid moves away and turns the „closed‟ form of the

lipase into an „open‟ form and allows the interaction between its hydrophobic internal

face and the hydrophobic residues that usually surround the lipase active center with

the substrate (Uppenberg et al., 1994; Carrasco-Lopez et al., 2009). When R. miehei

lipase was crystallized in the presence of the inhibitor n-hexylphosphonate ethyl ester,

the structure was in the open form (Brzozowski et al., 1991). True lipases display an

enhanced content of short, non-polar residues (usually valine, leucine and isoleucine),

which cluster at the protein hemisphere where the active site is located. These

residues facilitate the lipase attachment to the hydrophobic substrate aggregate, which

is then followed by the structural rearrangements responsible for the opening of the

lid (Fojan et al., 2000; Fernández-Lorente et al., 2007). Lipases from Bacillus subtilis

lacking the lid, and guinea-pig pancreatic lipase which features a “mini-lid” do not

undergo interfacial activation (Hjorth et al., 1993; Lesuisse et al., 1993).

Protein engineering of lipases

In view of wide application of lipases, difficulties in purifying their various

isoenzymes with overlapping biochemical properties and obtaining these enzymes in

bulk, attempts have been made to directly clone the genes encoding them. Beside

rather classical strategies such as immobilization, additives or process engineering,

molecular biology techniques nowadays represent probably the most important

methodology to tailor-design the enzyme for a given process. As described and

reviewed by Bornscheuer (2002a) protein engineering can be divided into three major

categories: functional expression of (iso-)enzymes, computer guided rational protein

Introduction

24

design and directed evolution. The first example of protein engineering of a lipase is

the work on Pseudomonas mendocina lipase (Gray et al., 1988). Bacterial lipases

from various Bacillus species are over-expressed in E. coli. However, many enzymes

which are used for a variety of biotransformations are not amenable to these systems

(Jaeger & Reetz, 1998). Pseudomonas species lipase require functional assistance of

~30 different cellular proteins before they can be recovered in an active state,

indicating that folding and secretion are highly specific processes (Rosenau & Jaeger,

2000). With the increasing knowledge of lipase structure and function, interest in

protein engineering of lipases is increasing. Some of the lipases, which are engineered

for their altered biochemical properties are listed (Table 1. 4).

The demand for the production of enantiometrially pure compounds has lead

to the directed evolution of highly enantioselective lipases (Reetz & Jaeger, 2003).

Bacterial lipases from P. aeruginosa and B. subtilis served as model enzymes to

demonstrate the potential of directed evolution. Variants of P. aeruginosa lipase with

high enantioselectivity towards both (S)- and (R)-2-methyldecanoic acid and p-

nitrophenylester has been created from a non-selective wild type enzyme (Liebeton et

al., 2000; Reetz et al., 2001; Zha et al., 2001). The solved crystal structures of P.

aeruginosa and B. subtilis lipases were used to rationalize amino acid exchanges

leading to increased enantioselectivity (Nardini et al., 2000; Van et al., 2001). A

comprehensive review on structural and biochemical data of C. rugosa lipolytic

isoenzymes (Lip1-Lip5), their biocatalytic reactivity and state-of-the-art of new

applications of wild-type and mutants of the wild-type has been presented

(Domínguez de María et al., 2006). C. antarctica lipase B is probably the most useful

lipase for numerous applications in organic synthesis. The thermostability, activity,

enantioselectivity and solubility of this enzyme have been improved by protein

engineering (Lutz, 2004). Site directed mutagenesis of Phe95

, Phe112

, Val206

and Val209

residues of Rhizopus delemar lipase led to a significant shift in the preference of the

mutant lipase for the hydrolysis of medium chain TAGs (Joerger & Haas, 1994). Pig

liver esterase, a prominent enantioselective enzyme, consists of several isoenzymes

with -, - and -subunits as the most dominant ones. The first functional over-

expression of an active -isoenzyme in the yeast P. pastoris was reported, allowing

the production of recombinant esterase at stable product quality without interfering

influences of other isoenzymes and hydrolases (Lange et al., 2001).

Introduction

25

Table 1. 4. Applications of engineered lipases.

Strain Application Effect of mutations Reference

Humicola

lanuginose

Detergent

additive

Increase in thermostability,

resistance to proteolytic degradation

Increase in specific activity

(Boel & Huge-Jensen,

1989)

(Andersen et al., 2011)

Pseudomonas

glumae

Detergent

additive

Resistance to oxidative degradation

Improvement of proteolytic

resistance

(Frenken et al., 1993)

(Batenburg et al.,

1991)

Pseudomonas

mendocina

Peracid

bleaching

systems

Alteration of substrate specificity

Improvement of perhydrolysis to

hydrolysis ratio

(Boston et al., 1997)

(Bott et al., 1994)

Candida

antarctica

Enantiomer

selectivity Alteration of substrate specificity (Engström et al., 2010)

Rhizopus

delemar Biocatalysis Alteration of chain length selectivity (Klein et al., 1997)

Geotrichum

candidum Biocatalysis Substrate specificity

(Holmquist et al.,

1997)

Application of lipases

Lipases are the most widely used class of enzymes in biotechnology with their

applications in organic synthesis and kinetic resolution of racemic compounds

(Schmid & Verger, 1998; Hasan et al., 2006). Three main reasons for their wide use

are:

a. Commercial preparations of many lipases are available. The major application is

in fat splitting. These enzymes become a convenient and preferred choice when

enzyme catalysis in low water media is developed.

b. Lipase reactions can be carried out in heterogeneous media, as a large number of

lipases show interfacial activation. Lipases have evolved usually stable structures

that may survive even the effect of organic solvents (Schmid & Verger, 1998).

c. Lipases have broad substrate specificities. Esters of fatty acids as well as alcohols

of various chain lengths are hydrolyzed. Similarly TAGs formed from fatty acids

of varying chain lengths are also hydrolyzed. In addition, lipases catalyze novel

reactions such as inter-esterification and esterification with enantioselectivity and

Introduction

26

regioselectivity (Sommer et al., 1997; Pandey et al., 1999).

The versatility of enzymatic properties of lipases in catalyzing different kinds

of reactions associated with their different species endows these enzymes with

important and vast application potential (Sharma et al., 2001; Yesiloglu & Baskurt,

2008). Lipases, like most specialty and industrial enzymes are increasingly produced

via recombinant DNA technology (Hasan et al., 2006). The substrate specificity,

chemo-, regio- and enantioselective biotransformation properties of lipases make

them excellent alternatives for classical chemical synthesis with industrial

applications. Table 1. 5 present a summary of the applications. The most common

applications of lipases are summarized below.

1. Food Industry

Lipases in food processing industry are used for the modification and

breakdown of biomaterials. Most of the commercial lipases produced are utilized for

flavor improvement in dairy products and processing of foods, such as meat,

vegetables, fruit, smoked carp, baked foods and beer (Freire & Castilho, 2008).

Immobilized R. miehei lipase is used extensively to carry out transesterification

reactions e.g., replacing palmitic acid in palm oil with stearic acid to produce the

desired stearic-oleic-stearic TAG (Sharma et al., 2001). Wheat, barley, corn and

canola seed lipases are used to produce low molecular weight esters in an organic

environment (Liaquat & Apenten, 2000). Lipases are generally used in the production

of a variety of products ranging from fruit juices to vegetable fermentation (Pandey et

al., 1999).

2. Detergent industry

An important application of lipases is that of an additive in industrial laundry

and household detergent formulations (Wiseman, 1995). The main features necessary

are stability under the conditions of washing (pH between 10.0 and 11.0 and

temperatures between 30 C and 60 C). Lipases function in the removal of stains

from fabrics (Aaslyng et al., 1991). They are also used in the synthesis of surfactants

for soaps, shampoos and dairy products (Pandey et al., 1999; Hasan et al., 2006). Rice

(Oryza sativa) phospholipase (Bhardwaj et al., 2001) and oat (Avena fatua) seed

lipases (Mohamed et al., 2000), which are stable at alkaline pH and temperatures of

about 60 C present suitable features for their use in detergents (Barros et al., 2010).

Introduction

27

Table 1. 5. Plant lipases and their biotechnological applications.

Lipase source Application Reference

Barley seed

(Hordeum vulgare)

Production of low molecular

weight esters

(Liaquat & Apenten,

2000)

Maize seed

(Zea mays)

Production of low molecular

weight esters

(Liaquat & Apenten,

2000)

Linseed

(Linum usitatissimum)

Production of low molecular

weight esters

(Liaquat & Apenten,

2000)

Rapeseed

(Brassica napus)

Production of low molecular

weight esters, Esterification

(Liaquat & Apenten,

2000)

Black-Cumin seeds

(Nigella sativa) Synthesis of structured lipids (Tuter et al., 2003)

Castor bean seed

(Phaseolus vulgaris) Synthesis of structured lipids (Tuter et al., 2003)

Wheat germ

(Triticum aestivum) Esterification (Xia et al., 2009)

Vernonia seed

(Vernonia galamensis) Hydrolysis of oils (Ncube & Read, 1995)

Papaya

(Carica papaya)

Fats and oils modification

Esterification and

interesterification

Asymmetric resolution of drugs

(Maria et al., 2006)

3. Oils and Fats

Lipase applications in the oil industry are enormous, as it reduces expenses

with energy and minimizes thermal degradation of compounds in comparison to

traditional chemical processes (Freire & Castilho, 2008). Interesterification and

hydrogenation properties are used in the preparation of glyceride products, which are

used in the production of margarine and butter. The partially purified lipase from

Nigella sativa seed was used to enrich borage oil with -linolenic acid (Tuter et al.,

2003). Hypolipidemic effects of blended oils and balanced fatty acid composition

obtained by interesterification of coconut oil and rice bran oil or sesame oil using R.

miehei lipase is reported (Reena & Lokesh, 2007).

4. Pharmaceutical industry

The ability of lipases to resolve racemic mixtures by the synthesis of a single

Introduction

28

enantiomer is currently exploited for drug production by the pharmaceutical industry

(Jaeger et al., 1999). Microbial lipases are extensively used to enrich polyunsaturated

fatty acids (PUFA) from animal and plant lipids. Free PUFA, their mono- and

diacylglycerides are used to produce a number of pharmaceuticals which are

anticholesterolemics, anti-inflammatories and thrombolytics (Gill & Valivety, 1997).

Pure (s)-ibuprofen, a non-steroidal anti-inflammatory drug was synthesized using

lipase catalyzed kinetic resolution via hydrolysis and esterification (Lee et al., 1995).

The biosynthesis of a model of flavor ester hexyl butyrate, by lipase catalyzed

chemical reactions under mild conditions is of commercial interest (Chang et al.,

2003).

5. Biodiesel

The use of lipase in biodiesel production has shown promising results.

Enzymatic transesterification reactions have been used to achieve higher yields in

biodiesel production, using refined oil compared to crude oils. The methanolysis of a

soybean oil and methanol using R. oryzae lipase in a solvent-free reaction system was

investigated. It is anticipated that such soybean oil methyl esters will be used as

biodiesel fuel (Kaieda et al., 2004). The oil-FFA, which can be used as the raw

material, is also completely converted into alkyl esters (Fukuda et al., 2001).

6. Pulp and paper industry

A unique enzymatic pitch control system has been developed to degrade

TAGs present in mechanical pulp slurry. Lipases produced from Aspergillus species,

C. rugosa and lipase powders produced from Candida cylindracea are used in pulp

and paper industry. Lipolytic enzymes are used to remove pitch, the lipid fraction of

wood that interferes with the elaboration of paper pulp. They also help in the removal

of lipid stains during paper recycling and to avoid the formation of sticky materials

(Hasan et al., 2006).

7. Biodegradable polymer production

Lipases are used as biocatalyst in the production of useful biodegradable

polyesters. 1-Butyl oleate is synthesized by direct esterification of butanol and oleic

acid to reduce the viscosity of biodiesel in winter use. The mixture of 2-ethyl-1-hexyl

esters is obtained in a good yield by enzymatic transesterification from rapeseed oil

fatty acids for use as a solvent. Y. lipolytica lipase was used as catalyst in the

Introduction

29

enzymatic ring-opening polymerization of ε-caprolactone (Barrera-Rivera et al.,

2012)

8. Cosmetics

Lipases have potential application in cosmetics and in aroma production

(Metzger & Bornscheuer, 2006). Mono- and diacyl glycerols are produced by

esterification of glycerols and are used as a surfactant in cosmetics and perfume

industries. The use of the lipase in place of the conventional acid catalyst enhances

product quality with minimum down streaming refining (Hasan et al., 2006).

9. Waste treatment

Lipases are utilized in activated sludge and other aerobic waste processes

where thin layers of fats are constantly removed from the surface of aerated tanks to

permit oxygen transport. Effective breakdown of solids clearing and prevention of fat

blockage or filming in waste systems are important in many industrial operations. For

e.g., sewage treatment, cleaning of holding tanks, septic tanks, grease traps and

degradation of organic debris (Rigo et al., 2008).

Rice (Oryza sativa)

Rice belongs to the family of grasses, Graminaeae (Poaceae). It is the most

important food cereal grown globally, serves as the staple food and supplies more

than 50 % of the required nutrients for a large section of the population (Sasaki &

Burr, 2000). The global production of rice has been estimated to be 650 million

tonnes. India stands first in rice area and second in rice production, after China

contributing 21.5 % of global rice production. More than 40,000 varieties of rice have

been reported worldwide. Approximately, 400 notified rice varieties are being

cultivated in India. Rice is a nutritious cereal crop used mainly for human

consumption and is the main source of energy (Table 1. 6) (Gopalan et al., 2007).

Rice bran, a by-product obtained during milling of rice has a colossal potential

for use in the food and feed industry. Bran constitutes 10 % of the rough grain and

composed of pericarp, seed coat, nucellus and aleurone layer (Figure 1. 7). The

chemical composition shows that its protein is of high nutritional value (Kennedy &

Burlingame, 2003). Bran is also a good source of Vitamin E and oryzanol, although

their concentrations vary substantially, depending on the origin (Nicolosi et al., 1993).

Bran is known to be rich in water soluble polysaccharides, proteins, lipids and

Introduction

30

vitamins (Prakash, 1996). Rice bran in the diet has shown to prevent different

diseases, including cancer, hyperlipidemia, fatty liver, hypercalciuria, kidney stones

and heart disease (Jariwalla, 2001). The United Nations Industrial Development

Organization has designated rice bran as an under-utilized raw material (Zhang et al.,

2009). Rice contains a significant quantity of fatty acid based energy storage

molecules, typically containing mono- or PUFA. These oil bodies serve as an energy

reserve during germination and radicle growth. During traditional milling of brown

rice, the TAGs in bran come in mutual contact with an endogenous lipase leading to

the release of FFA. The high concentration of endogenous TAGs and the released

FFA are thus sensitive to oxidative degradation leading to rancidity. Complete

utilization of rice bran is therefore hampered (Funatsu. et al., 1971).

Table 1. 6. Composition per 100 g of edible portion of milled rice.

Nutrient Quantity Nutrient Quantitiy

Calories (kcal) 345.0 Calcium (mg) 10.0

Moisture (g) 13.7 Iron (mg) 0.7

Carbohydrates (g) 78.2 Magnesium (mg) 90.0

Protein (g) 6.8 Riboflavin (mg) 0.06

Fat (g) 0.5 Thiamine (mg) 0.06

Fibre (g) 0.2 Niacin (mg) 1.9

Phosphorous (mg) 160.0 Folic acid (mg) 8.0

Minerals (g) 0.6 Copper (mg) 0.14

Adapted from Gopalan et al., 2007

Figure 1. 7. The parts of a rice grain

Introduction

31

The inhibition or suppression of rice bran lipase is a major concern of the rice

industry. Attempts have been made to chemically inactivate lipase to obtain better

quality oil and bran fit for food and feed purposes. Several physical and chemical

methods have been evolved to inactivate lipase and stabilize rice bran, however with

limited success. These methods include microwave heating (Lakkakula et al., 2004;

Zigoneanu et al., 2008), ohmic heating (Ramezanzadeh et al., 2000; Loypimai et al.,

2009), extrusion (Zhu & Yao, 2006), chemical treatment using HCl (Nasirullah et al.,

1989), refrigeration and lowering pH (Amarasinghe et al., 2009). Heat treatment is the

most common method to stabilize rice bran. Temperatures above 120 C are used to

denature the enzyme responsible for lipid degradation. The extrusion (dry heating)

cookers have been ideal for stabilization because excess moisture is not added

eliminating the need for drying (Orthoefer, 2005).

Rice bran lipase

Most of the lipolytic activity in rice is found in the bran (Borgston &

Brockman, 1984). The isolation and purification of a lipase was first reported by

Funatsu et al., (1971) and Shastry & Rao (1971). The lipase was purified by

ammonium sulfate precipitation followed by a series of chromatographies using

DEAE cellulose, Sephadex-G75 and CM-Sephadex C-50 in the presence of Ca2+

ion.

The specific activity of the purified enzyme was 4.7 units/mg protein. The molecular

mass determined was found to be 40,000 Da with two subunits. Isoelectric focusing of

the homogeneous protein indicated the isoelectric point of 8.56. It preferentially

cleaved fatty acids from the sn1 and sn3 positions of TAGs (Funatsu et al., 1971). The

physico-chemical properties of the enzyme were reported by Aizono et. al., (1973).

The enzymatic properties with reference to pH, temperature, effect of Ca2+

ions,

EDTA and substrate specificity were also reported (Aizono et al., 1973). Shastry and

Rao (1971) reported the purification of rice bran lipase by ammonium sulphate

precipitation and ion exchange chromatography. The enzyme showed higher activity

towards tributyrin as compared to tripalmitin and exhibited a molecular weight of

40,000 Da.

Purification and characterization of rice bran lipase II (Lipase-II) with a

molecular weight of a 33,300 Da was reported (Aizono et al., 1976). The enzyme

showed the optimum pH between 7.5 and 8.0, and the optimum temperature at about

27 C. It was stable over the pH range from 5 to 9.5 and below 30 C. The enzyme

Introduction

32

exhibited a high specificity toward TAGs having short carbon chain fatty acids,

although it was capable of hydrolyzing the ester bonds in the olive oil. The

comparison of the properties of lipase isolated by Funatzu et al., (1971) showed

marked differences to that isolated by (Aizono et al., 1976). Therefore, these were

designated as Lipase-I and –II. Rajeshwara & Prakash (1995) purified and

characterized a lipase from rice bran with higher yields and activity. The enzyme

showed an UV absorption maximum of 276 nm with a pH optimum of 7.5 and

temperature optimum of 30 C. The molecular weight of enzyme was 30,000 Da and

found to contain 16 % -helix, 39 % -sheets and the rest being random coil. A

thermally stable phospholipase A2 of 9,400 Da from rice bran has been purified to

homogeneity and characterized. The enzyme was found to be a glycoprotein and

showed maximum activity at 80 C and pH 11.0. It preferentially hydrolyzed the sn2

position of phosphatidylcholine, but apparently exhibited no positional specificity

toward TAG (Bhardwaj et al., 2001). Lipase-II was reported to be the major lipase in

rice bran (Aizono et al., 1976; Fujiki et al., 1978; Rajeshwara & Prakash, 1995).

Heterologous expression and functional properties of various prokaryotic and

eukaryotic lipases have been documented (Aloulou et al., 2006; Yu et al., 2007;

Larsen et al., 2008; Sabri et al., 2009 ). In contrast, remarkably little is reported on the

molecular cloning and functional expression of cereal lipases, despite their

fundamental importance. A lipase gene from rice (Oryza sativa cv. Dongjin) has been

cloned and expressed in E. coli. The expressed protein was about 40,000 Da,

exhibited specificity to tributyrin (Kim, 2004). Recently, an esterase from rice bran

was identified, cloned and successfully expressed as a recombinant protein, which

although contained the conserved esterase/lipase motif, the essential active site serine

(GXSXG) was replaced by cysteine (Chuang et al., 2011).

Introduction

33

Aim and scope of the present investigation

Rice (Oryza sativa), one of the world‟s important cereals is the staple food for

a large section of the population and supplies more than 50 % of the required protein

for this group. Rice bran, the separated brown layer, which contains the pericarp and

germ, a by-product obtained during the abrasive milling of brown rice is enriched in

protein, lipid, vitamins, minerals and water-soluble polysaccharides. Rice bran has

enormous potential for exploitation and utilization in foodstuffs, medicines,

healthcare and animal feeds (Prakash, 1996). This by-product with tremendous

potential as a rich source of nutrients is under-utilized. The rapid deterioration and

development of off flavours due to the released fatty acids, catalyzed by a lipase

limits the use of rice bran, both as food and feed. A lipase, is chiefly responsible for

development of off-flavours after processing. During traditional milling of brown rice,

the TAGs in bran come in mutual contact with lipase. This results in the development

of FFA, which makes rice bran unpalatable. The inhibition or suppression of rice bran

lipase is a major concern of the rice industry. Attempts have been made to chemically

inactivate lipase to obtain better quality oil and bran fit for food and feed purposes.

Several physical and chemical methods have been evolved to inactivate rice bran

lipase however with limited success (Jiaxun, 2001; Sharma et al., 2004; Raghavendra

et al, 2007).

Newer scientific approaches to control lipase activity in rice have to be based

on an understanding of the structure function relationship of rice bran lipase at the

molecular level. Therefore, the starting point for developing an efficient method for

stabilizing rice bran would be a biotechnological approach. In an attempt to correlate

the role of rice bran lipase at the molecular level for developing efficient methods to

restrict its activity, the major focus of this study is the molecular cloning and

functional expression of the lipase. The main objectives of the present investigation

are:

Molecular cloning of the lipase gene from rice (Oryza sativa cv. Indica-IR 64).

Expression in E. coli and Pichia pastoris and characterization of the lipase.

Temporal expression profiling of lipase during seed development and

germination.

It is expected that these studies will lead to a better understanding of the

structure-function relationship of the Lipase-II designated as rice bran lipase (RBL) in

Introduction

34

this study. A molecular approach, in combination with the functional analysis of

lipase would provide a deeper insight into physiological role during seed germination,

growth and development of rice caryopsis. The molecular level studies would

eventually provide a platform for the design of a tailored lipase to prevent enzymatic

rancidity.