15

Review of

Literature

Bacterial

Lipase

Lipase

Lipase

Bacterial

16

17

2. Review of Literature

Industrial biotechnology, also known as white biotechnology, is the application of

modern biotechnology to the sustainable production of chemicals, materials, and fuels from

renewable sources, using living cells and/or their enzymes. This field is widely regarded as

the third wave of biotechnology, distinct from the first two waves (medical or red

biotechnology and agricultural or green biotechnology). Much interest has been generated

in this field mainly because industrial biotechnology is often associated with reduced

energy consumption, greenhouse gas emissions, and waste generation, and also may enable

the paradigm shift from fossil fuel-based to bio-based production of value added chemicals.

Industrial biotechnology is a rapidly growing field which involves the use of enzymes and

microorganisms. Enzymes have been used by men since biblical times either as vegetables

rich in enzymes or as microorganisms and their products (in brewing processes, baking

and production of alcohol). Beer and yoghurt also owe their flavor and texture to a range of

enzyme- producing organisms that were domesticated many years ago. Enzymes are

natural catalysts which play a diversified role in many aspects of everyday life.

2. 1 Hydrolases

The hydrolase family includes a group of enzymes that catalyze bond cleavage by

reacting with water. Amongst others, these consist of lipases, proteases, amidases, epoxide

hydrolases, nitrilases, and glycosidases. Hydrolases are placed in Class 3 according to the

IUB classification of enzymes by the Enzyme Commission, and these are further classified

by the type of bond hydrolyzed; for instance, lipases are classified as EC 3.1.1.3 as they

hydrolyze the carboxyl ester bonds of triacylglycerols. Currently, the biotechnological

applications of hydrolases are of special interest as they have some advantageous

characteristics which make them ideally suited for industrial use. Strikingly, most of the

enzymes used in industry are microbial enzymes, originating either from bacteria, fungi

and yeasts.

18

2.2 Lipases

Lipase (triacylglycerol acylhydrolase, EC 3.1.1.3) catalyzes the hydrolysis of the

carboxyl ester bonds in triacylglycerols to produce diacylglycerols, monoacylglycerols,

fatty acids and glycerol. In addition, lipases catalyze the hydrolysis and transesterification

of other esters as well as the synthesis of esters. Many lipases exhibit enantioselective

properties.

Figure 2.1 Hydrolysis or synthesis (acylglycerols/esters) reactions catalyzed by lipases

The literature pertaining to lipases has been reviewed by several investigators

(Gupta et al., 2004; Hasan et al., 2006; Hasan et al., 2009; Jaeger and Eggert, 2004; Kapoor

et al., 2012; Salihua et al., 2012). First lipase was discovered in pancreatic juice in the year

1856 by Claude Bernard. Animal pancreatic extracts were traditionally used as the source

of lipase for commercial applications. However, microbial sources of lipase were explored

when industrial potential of lipases enhanced and their demand could not be met by the

supply from animal sources. The number of available lipases has increased mainly as a

R’-OH

19

result of achievements made in the cloning and expression of enzymes from

microorganisms, as well as of an increasing demand for these biocatalysts with novel and

specific properties such as specificity, stability, pH, and temperature (Bornscheuer et al.,

2002; Menoncin et al., 2009).

Lipases are widely distributed in animal, plants and microorganisms (Bornscheuer,

2002), however, microbial lipases are commercially most important mainly because of the

ease of their cultivation and genetic manipulation to obtain higher yield (Hasan et al.,

2006). Commercially important microbial lipases are produced from bacteria, fungi and

yeast (Abada, 2008; Babu and Rap, 2007). Some lipase-producing microorganisms are listed

in Table 2.1.

The industrial demand for new sources of lipases with different catalytic

characteristics stimulated the isolation and selection of new strains. 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 among

others (Sharma et al., 2001).

2.2.1 Bacterial lipases

Among the bacterial lipases, the enzymes produced from Bacillus sp. possess

properties that make them potential candidates for biotechnological applications. The most

common lipase-producing bacterial strains are Bacillus subtilis, Bacillus pumilus, Bacillus

licheniformis, Bacillus coagulans, Bacillus stearothermophilus and Bacillus alcalophilus. In

addition, Pseudomonas sp., Pseudomonas aeruginosa, Burkholderia multivorans,

Burkholderia cepacia, and Staphylococcus caseolyticus were also reported as lipase

producers. Ertugrul and co-workers (2007) isolated 17 bacterial strains that could grow

and produce lipase on media based on olive oil mill waste. In these strains, maximum

intracellular lipase activity was found to be 168 UmL−1 after medium optimization. Shariff

et al. (2007) isolated a thermophilic bacterium, Bacillus sp. strain L2, from a hot spring in

Perak, Malaysia. An extracellular thermostable lipase activity was detected through plate

and broth assays at 70 °C after 28 h of fermentation.

Kiran and co-workers (2008) isolated 57 heterotrophic bacteria from the marine

sponge Dendrodoris nigra, of which 37 % produced a clear halo around the colonies on

20

tributyrin agar plates for lipase production. The strain Pseudomonas MSI057 produced the

largest zones on tributyrin agar plate and the lipase activity was 750 UmL-1 after

optimization. Carvalho et al. (2008) isolated a bacterial strain from petroleum-

contaminated soil and designated it as Biopetro-4. Abada (2008) produced lipase from

Bacillus stearothermophilus AB-1 isolated from air. Takaç and Marul (2008) isolated

microbial cultures from soil enriched by periodic sub-culturing of samples in nutrient broth

containing 1% (v/v) tributyrin. The isolation process was performed on tributyrin agar

(TBA) plates and Bacillus species producing large zone on TBA were selected. Active

colonies were re-streaked on TBA agar for purification.

Bora and Bora (2012) isolated an extracellular alkaline lipase-producing bacterial

strain Bacillus sp. LBN2 from soil sample of hot spring of Arunachal Pradesh, India. The

cells were cultivated in a mineral medium with maximum production at 1 % groundnut oil.

The optimum temperature and initial medium pH for lipase production by the organism

were 50 C and 9.0 respectively.

Bacterial lipases may be intracellular, membrane-bound or extracellular. Lee and

Park (2008) reported the production of intracellular lipase from a strain of Bacillus

clausii which could grow only on glycerol and simple lipids but not on long chain

triglycerides. Ertugrul et al. (2007) observed the production of both intracellular and

extracellular lipase in Bacillus sp. Boekema et al. (2007) documented the production of

extracellular lipase as a consequence of secretion of accumulated intracellular lipase by

membrane-bound chaperones.

Kiran and co-workers (2008) isolated 57 heterotrophic bacteria from the marine

sponge Dendrodoris nigra, of which 37 % produced a clear halo around the colonies on

tributyrin agar plates for lipase production. The strain Pseudomonas MSI057 produced the

largest zones on tributyrin agar plate and the lipase activity was 750 UmL-1 after

optimization. Carvalho et al. (2008) isolated a bacterial strain from petroleum-

contaminated soil and designated it as Biopetro-4. Abada (2008) produced lipase from

Bacillus stearothermophilus AB-1 isolated from air. Takaç and Marul (2008) isolated

microbial cultures from soil enriched by periodic sub-culturing of samples in nutrient broth

containing 1% (v/v) tributyrin.

21

Table 2.1 List of some lipase-producing microorganisms

Sources: Gupta et al., 2004; Treihel et al., 2010

Microorganisms References

Bacterial

Acinetobacter calcoaceticus Dharmsthiti et al. (1998); Jaeger et al. (1999); Pandey et al. (1999)

Acinetobacter radioresistens Liu and Tsai (2003)

Acinetobacter sp. Snellman et al. (2002)

Acinetobacter sp. Barbaro et al. (2001)

Aeromonas caviae AU04 Velu et al. (2012)

Arthrobacter sp. Pandey et al. (1999)

Bacillus alcalophilus Ghanem et al. (2000)

Bacillus atrophaeus Bradoo et al. (1999)

Bacillus pumilus Jaeger et al. (1999) Bacillus stearothermophilus Bradoo et al. (1999); Jaeger et al. (1999)

Bacillus subtilis Jaeger et al. (1999); Ruiz et al. (2005)

Bacillus thermocatenulatus Jaeger et al. (1999); Pandey et al. (1999)

Bacillus sp. Nawani and Kaur (2000); Bora and Bora (2012)

Bacillus thermoleovorans Rua et al. (1997)

Chromobacterium viscosum Taipa et al. (1995)

Lactobacillus plantarum Lopes et al. (1999)

Pseudomonas fluorescens Kojima et al (1994)

Pseudomonas sp. Sarkar et al. (1998)

Staphylococcus aureus Gotz et al. (1998) Staphylococcus aureus Sarkar et al. 2012

Staphylococcus xylosus Mosbah et al. (2005)

Stenotrophomonas maltophilia Hasan-Beikdashti et al. (2012)

Thermosyntropha lipolytica Gumerov et al. 2012

Fungal and Yeast

Candida cylindracea Muralidhar et al. (2001)

Candida rugosa Rajendran et al. (2008); Zhao et al. (2008)

Candida utilis Grbavcic et al. (2007)

Fusarium solani Knight et al. ( 2000)

Penicillium cyclopium Chahinian et al. (2000) Pichia burtonii Sugihara et al. (1995)

Rhizopus sp. Macedo et al. (2003)

Rhodotorula mucilaginosa Potumarthi et al. (2008)

Trichosporon asahii Kumar and Gupta (2008)

Yarrowia lipolytica Alonso et al. (2005); Fickers et al. (2006); Amaral et al. (2007); Kar et al. (2008); Lopes et al. (2009)

22

The isolation process was performed on tributyrin agar (TBA) plates and Bacillus species

producing large zone on TBA were selected. Active colonies were re-streaked on TBA agar

for purification.

Bora and Bora (2012) isolated an extracellular alkaline lipase-producing bacterial

strain Bacillus sp. LBN2 from soil sample of hot spring of Arunachal Pradesh, India. The

cells were cultivated in a mineral medium with maximum production at 1 % groundnut oil.

The optimum temperature and initial medium pH for lipase production by the organism

were 50 C and 9.0 respectively.

Bacterial lipases may be intracellular, membrane-bound or extracellular. Lee and

Park (2008) reported the production of intracellular lipase from a strain of Bacillus

clausii which could grow only on glycerol and simple lipids but not on long chain

triglycerides. Ertugrul et al. (2007) observed the production of both intracellular and

extracellular lipase in Bacillus sp. Boekema et al. (2007) documented the production of

extracellular lipase as a consequence of secretion of accumulated intracellular lipase by

membrane-bound chaperones.

2.2.1.1 Classification of bacterial lipases

Lipases belong to the family of serine hydrolases and their activity relies on a

catalytic triad comprising of serine, histidine, and aspartate and α/β hydrolase fold.

Bacterial lipolytic enzymes were classified into 8 families and the largest family was

subdivided into 6 sub-families by Arpigny and Jaeger (1999) based on the conserved

sequence motifs and biological properties of the enzymes. Family I comprise most of the

lipases produced by Pseudomonas, Bacillus and Staphylococcus. True lipases belong to this

family. These lipases possess the conventional catalytic pentapeptide Gly-X-Ser-X-Gly.

Family II lipases exhibit Gly-Asp-Ser-Leu motif at the active site. Esterases produced

by Streptomyces, Aeromonas and Salmonella belong to this family. Family III comprise

lipases of Streptomyces sp. but unlike family II esterases these are extracellular lipases.

Lipases which display similarity with mammalian hormone sensitive lipases are grouped

under Family IV while lipases of mesophilic bacteria like Pseudomonas

oleovorans and Haemophilus influenza belong to family V. Family VI lipases are the smallest

esterases and the active enzymes are dimeric. Family VII lipases are large esterases and

23

their amino acid sequence is homologous to that of eukaryotic acetyl choline esterases.

Family VIII lipases are similar to β-lactamases. The sequences of few other enzymes could

not be grouped into any of the eight super families described by Arpigny and Jaeger (1999)

and have been arbitrarily classified as new family 9 and 10. A cold active lipase reported

by De Pascale et al. (2008) could not fit into the traditional classification and hence

reported as a lipase belonging to a novel lipolytic family.

MELDB is another comprehensive database of microbial lipases and esterases

(Kang et al., 2009). The orphaned lipases which do not belong to any of the eight

superfamilies but arbitrarily grouped with them in the traditional classification have been

out-grouped in the MELDB database. This classification was done with conserved

sequences of enzymes based on a local sequence alignment and a graph clustering

algorithm (Tribe MCL).

2.2.2 Fungal Lipases

The first work on fungal lipases was reported by Ghosh et al. (1996). In 1994, Novo

Nordisk introduced the first commercial recombinant lipase ‘Lipolase’ which originated

from the fungus Thermomyces lanuginosus and was expressed in Aspergillus oryzae. Most of

the commercially important lipase-producing fungi belong to the genera Rhizopus sp.,

Aspergillus sp., Penicillium sp., Geotrichum sp., Mucor sp., and Rhizomucor sp. Colen et al.

(2006) isolated 59 lipase-producing fungal strains from Brazilian savanna soil using

enrichment culture techniques. An agar plate medium containing bile salts and olive oil

emulsion was employed for isolating and growing fungi in primary screening assay.

Twenty one strains were selected by the ratio of the lipolytic halo radius and the colony

radius. Eleven strains were considered and among them, the strain identified as

Colletotrichum gloesporioides was the most productive. In another work, Cihangir and

Sarikaya (2004) isolated a strain of Aspergillus sp. from soil samples of different regions of

Turkey having lipase activity of 17 UmL−1. Vargas et al. (2008) studied the production of

lipase by Penicillium simplicissimum and obtained an activity of 30 U gds−1. Both Penicillium

verrucosum and Penicillium simplicissimum were isolated from the babassu oil industry.

24

2.2.3 Yeast Lipases

According to Vakhlu and Kour (2006), the main terrestrial lipase- producing species

of yeasts are Candida rugosa, Candida tropicalis, Candida antarctica, Candida cylindracea,

Yarrowia lipolytica, Rhodotorula glutinis, and Pichia burtonii. The genes that encode lipase

in Candida sp., Geotrichum sp., Trichosporon sp., and Y. lipolytica were cloned and over-

expressed (Wang et al., 2007).

Potumarthi et al. (2008) collected marine soil samples from the surroundings of an

oil extraction platform in the Arabian Sea and isolated colonies, which were transferred to

the petriplates containing 2 % tributyrin and incubated at 35 °C for 3–4 days. The colonies

showing the largest hydrolysis halos zone were selected. The most effective strain for

lipase production was identified as Rhodotorula mucilaginosa (MTCC 8737) by its

phenotypic characteristics.

Kumar and Gupta (2008) obtained 15 yeast isolates from petroleum and oil sludge

areas in Delhi, India. The isolates were purified and checked for their lipolytic potential.

Among these yeast strains, one strain was selected for further studies, based on the largest

halo of lipolysis. On the basis of sequence homology, this strain belonged to Trichosporon

asahii genus and shared 99 % homology with the already existing database. The

microorganisms isolated from this oil included several strains of lipase-producing yeasts

which were identified as Saccharomyces cerevisiae, Candida wickerhamii, Williopsis

californica, and Candida boidinii. The lipase activity was noted to be intracellular in

Saccharomyces cerevisiae. The three-phase olive oil extraction process generated a dark-

colored effluent, usually termed olive oil mill wastewater.

D’annibale et al. (2006 b) investigated the valorization of oil mill waste water by

using it as a possible growth medium for the microbial production of extracellular lipase.

Among the 12 strains tested, the most promising strain was Candida cylindracea. The most

potential lipase producer from yeasts reported in the literature is Candida sp.

2.2.4 Lipases from extremophiles

Extremophiles are the organisms able to survive under extremes conditions of

temperature, pressure, low water activity, salinity, acidity, alkalinity, radiation etc. As a

result, extremophiles have the potential to produce uniquely valuable biocatalysts that

25

function under conditions in which, their usually non-extremophilic counterparts could

not.

2.2.4.1 Lipases from psychrophiles

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 bacterium 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 produced lipases

that possessed high activity, but not optimum, in the temperature range of 0 to 20 °C

(Feller et al., 1990). Consequently, genes for three lipases from Moraxella TA144 were

sequenced and cloned in E. coli (Feller et al., 1990). This was followed by isolation of many

other psychrophilic lipolytic bacteria including Acinetobacter calcoacetius LP009

(Pratuangdejkul and Dharmsthiti, 2000), Psychrobacter okhotskensis (Yumoto et al., 2003)

and the psychrotolerant bacterium Corynebacterium paurometabolum MTCC 6841 (Joshi et

al., 2006).

2.2.4.2 Lipases from thermophiles

Thermophiles are the most investigated extremophiles (Wiegel et al., 1998). These

enzymes are generally the most stable at high temperature and stable in organic solvents

(Ejima, et al., 2004; Fucinos et al., 2005; Li and Zhang, 2005). Although there are some

enzymes from mesophilic sources that withstand elevated temperatures but such cases are

rare. 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 (Adams and Kelly, 1998; Eichler, 2001).

2.2.5 Commercial lipases and their industrial suppliers

Microbial lipases are industrially important enzymes. A number of commercial

products have been launched successfully worldwide. In 1994, Novo Nordisk introduced

26

the first commercial recombinant lipase ‘Lipolase’ from the fungus Thermomyces

lanuginosus and was expressed in Aspergillus oryzae. In 1995, two bacterial lipases were

introduced – ‘Lumafast’ from Pseudomonas mendocina and ‘Lipomax’ from Pseudomonas

alcaligenes by Genencor International (Jaeger and Reetz, 1998). Table 2.2 enlists the

names of various suppliers of commercial lipases.

Table 2.2. Suppliers of commercial lipases

Sources: Modified from Gupta et al., 2004; Hasan et al., 2006

2.3 Screening of lipase- producing microorganisms

Lipolysis could be detected directly by changes in the appearance of the substrate

such as tributyrin and triolein, which were emulsified mechanically in various growth

media and poured into petridishes. Lipase production was indicated by the formation of

clear halos around the colonies grown on tributyrin containing agar plates (Ertugrul et al.,

Commercial lipase Source Supplier Application

Lumafast Pseudomonas menodocina

Genencor , USA Detergent

Lipomax Pseudomonas alcaligenes

Genencor International, USA

Detergent

Lipofast NA Advanced Biochemicals, India

Detergent

Lipase AH Pseudomonas cepacia

Amano Pharmaceuticals, Japan

Organic synthesis

Lipase K-10

Pseudomonas sp.

Amano Pharmaceuticals, Japan

Organic synthesis

Amano P, P-30, PS, LPL-80, LPL-200S

Pseudomonas cepacia

Amano Pharmaceuticals, Japan

Organic synthesis

Lipase 50P

C. viscosum

Biocatalysts, UK

Biotransformations, chemicals

Combizyme 61P (proteinase/lipase mix)

NA Biocatalysts, UK

Waste treatment

Greasex (lipase)

NA Novo Nordisk

Leather

27

2007; Jaeger et al., 1994; Kim et al., 2001). Bacillus strains were screened for lipolytic

activity on agar plates containing tributyrin or Tween 20 or Tween 80 (1%, w/v) and 2%

agar-agar (Fakhreddine et al., 1998; Zinterhofer et al., 1973). Lipolytic Bacillus sp LBN 4

was isolated on tributyrate agar medium using glycerol tributyrate as substrate (Bora and

Kalita, 2007).

Lipolytic activity on solid media could be visualized by using dyes such as Victoria

blue B, Spirit blue, Nile blue sulfate and Night blue (Shelley et al., 1987a). The drop in pH

due to the fatty acids released as a result of hydrolysis was observed by change in the

colour of indicators used (Scholze et al., 1999). There was a linear relationship between the

diameter of the fatty acid diffusion spot and the logarithm of the enzyme concentration.

This technique is very convenient for rapid screening of lipolytic microorganisms but

acidification of the medium due to the generation of acidic metabolism other than free fatty

acids, which are released by microbial lipases, can give false results. The fluorescent dye

Rhodamine B could also be used in plate assay containing emulsified olive oil to detect

lipolytic organisms where substrate hydrolysis caused the formation of orange fluorescent

halos around bacterial colonies visible upon UV irradiation and the lipase activities ranged

from 1 to 30 nkat (Kim et al., 2001; Kouker and Jaeger, 1987).

2.4 Lipase production and parametric optimization

Enzymes can be produced by submerged fermentation (SmF) or by a solid state

fermentation (SSF). SmF, which involves the growth of the microorganism as a suspension

in nutrient enriched liquid medium, is the preferred method for production of commercial

enzymes, principally because sterilization and process control are easier to engineer in

these systems. On the other hand, SSF is the growth of microorganisms on most substrates

in the absence of free-flowing water. The advantages of SSF processes over liquid batch

fermentation include smaller volumes of liquid required for product recovery, cheap

substrate, low cultivation cost for fermentation, and lower risk of contamination. Fungal

species are preferably cultivated in SSF, while bacteria and yeast are cultivated in SmF

(Dutra et al., 2008). Lipase production in SmF has been reported using batch, repeated-

batch, fed-batch and continuous fermentation (Treichel et al., 2009).

28

Many studies have been undertaken to define the optimal culture and nutritional

requirements for lipase production by submerged culture. Microbial lipases production is

greatly influenced by medium composition besides physicochemical factors such as

inoculum size and age, temperature, pH, dissolved oxygen concentration etc. (Aires-Barros

et al., 1994; Bora and Bora, 2012; Brune and Gotz, 1992; Jaeger et al., 1994; Kim et al.,

1996). The type and concentration of carbon and nitrogen sources influence lipase

production (Elibol and Ozer, 2001). The major factor for lipase activity has always been

reported as the carbon source, since lipases are inducible enzymes. Lipidic carbon sources

seem to be essential for obtaining a high lipase yield; however, a few authors have

produced good yields in the absence of fats and oils. These enzymes are generally produced

in the presence of a lipid such as oil or any other inducer, such as diacylglycerols, fatty

acids, hydrolysable esters, tweens, bile salts, and glycerol (Ghosh et al., 1996; Gupta et al.,

2004; Rathi et al., 2001; Sharma et al., 2001).

Nitrogen sources and essential micronutrients should also be carefully considered

for growth and production optimization. These nutritional requirements for microbial

growth are fulfilled by several alternative media as those based on defined compounds like

sugars, oils, and complex components such as peptone, yeast extract, malt extract media,

and also agro-industrial residues containing all the components necessary for

microorganism development. Generally, high productivity has been achieved by culture

medium optimization. An improper optimization of these factors leads to a lower

production of the enzyme. Fermentation conditions for enzyme production can be

optimized either by considering one variable at a time approach or by statistical approach.

2.4.1 Parametric optimization using one variable approach

2.4.1.1 Effect of inoculum size and age

The preliminary requirement for mass production of microbial cells as well as

enzymes involves determination of suitable inoculum size and age. Initially, microbial cell

growth as well as enzyme activity increase with increase in inoculum size, but declines

after a certain limit. The effect of inoculum size on lipase activity can be correlated with

total dissolved oxygen in the medium. Any variation in inoculum size from the optimum

29

concentration results in reduced enzyme yield. Generally 1-10 % (v/v) inoculum of

bacterial culture is sufficient for SmF reactions. Lee et al. (1999) reported that 1%

inoculum size was optimum for lipase production by Bacillus thermoleovorans ID-1 while

Hasan et al. (2006) reported 5 % in case of Bacillus sp. FH5. The maximum lipase yield

reached up to 251.78 U/ml by G. stearothermophilus B-78 at an inoculum size of 2.5 ml per

20 ml (Bayoumi et al., 2007).

A survey of the literature revealed that 12-24 h old inoculum would be best suited

for fermentation reactions of majority of microbial strains. A higher inoculum age is not

preferred at the industrial level. An inoculum of 18 h was found to be optimum for lipase

production by Thermoactinomyces vulgaris (Elwan et al., 1978) whereas 6 h and 24 h old

culture led to the maximum lipase production for Bacillus sp. (Hasan et al., 2003; Sidhu et

al., 1998b)

2.4.1.2 Effects of incubation period, temperature and pH

Incubation period is the time taken by the inoculated culture for synthesis of the

desired product (enzyme) utilizing the medium nutrients. However, the duration of

incubation depends up on the type of fermentation. Generally, the incubation period

required for production under SSF is higher as compared to SmF. Further, duration of

incubation also varies according to the type of microorganism used. The optimum period

for bacterial lipase production in SmF was reported to vary from 18 to 48 h (Ebrahimpour

et al., 2008; El-Shafei and Rezkallah, 1997; Handelsman and Shoham, 1994; Hasan et al.,

2006). Fungal strains and yeast strain require more incubation time as compared to

bacterial strain. Candida rugosa (Song et al., 2001) produced maximal titre of lipase after

60 h of incubation whereas Rhizopus oryzae (Hiol et al., 2000) and Penicillium wortmanii

(Costa and Peralta., 1999) required 4 and 5 days of incubation respectively.

Temperature is an important physical parameter affecting the production of enzyme

from a given microbial culture. Different types of microorganisms require different

optimum fermentation temperature for maximum enzyme production. Some

microorganisms such as Bacillus cereus (Dutta and Ray, 2009), Acinetobacter radioresistens

(Chen et al., 1998), Enterobacter agglomerans (Zhen-qian and Chun-yun, 2009) and

Pseudomonas sp. 7323 (Zhang and Zeng, 2008) showed 30 °C as the optimum temperature

30

for growth and lipase production. Hasan et al. (2006) reported the optimum temperature

of 37 °C for lipase production by Bacillus sp. FH5.

The pH of the growth medium is very important factor for microbial growth and

enzyme production. Depending upon the pH requirement, a microorganism may be

acidophilic, alkalophillic or both. Small variation from the optimum pH may lead to

significant drop in growth and enzyme production. Largely, bacteria prefer pH around 7.0

for best growth and lipase production, such as in case of Bacillus sp. (Sugihara et al., 1991),

Acinetobacter sp. (Barbaro et al., 2001) and Burkholderia sp. (Rathi et al., 2001). On the

other hand, bacterial lipases were reported to prefer alkaline pH (>7.0) for growth and

enzyme production (Dong et al., 1999; Sharma et al., 2002b; Wang et al., 1995). The

incubation period, temperature and pH for some of the lipolytic microorganisms are

presented in Table 2.3.

2.4.1.3 Effects of carbon and nitrogen sources on lipase production

The major factor for the expression of lipase activity has always been carbon source.

Sugihara et al. (1991) reported lipase production from Bacillus sp. in the presence of 1 %

olive oil in the culture medium whereas modest enzyme activity was observed in the

absence of olive oil even after prolonged cultivation. Production of lipase can be

significantly influenced by other carbon sources such as sugars, sugar alcohol,

polysaccharides, whey, casamino acids and other complex sources (Dharmsthiti and

Kuhasuntisuk, 1998; Gilbert et al., 1991a; Lotrakul and Dharmsthiti, 1997; Rashid et al.,

2001).

However, lipases from Pseudomonas aeruginosa EF2 (Gilbert et al., 1991a) and

Acinetobacter calcoaceticus (Mahler et al., 2000) were repressed in the presence of long-

chain fatty acids, such as oleic acid. Kanwar et al. (2002) reported the production of a

Pseudomonas sp. G6 lipase in the presence of n-alkane substrates, with a maximum

production of about 25 U/ml when n-hexadecane was the sole carbon source. Olive oil and

n-hexadecane were employed as the carbon source for producing an alkaline lipase from

Acinetobacter radioresistens (Liu and Tsai, 2003). A thermophilic Bacillus strain A30-1

(ATCC 53841) produced maximal levels of thermostable alkaline lipase when corn oil and

olive oil (1 %) were used as carbon source (Wang et al., 1995).

31

Table 2.3 Effect of incubation period, pH and temperature on lipase production

Sources: Modified from Hasan et al., 2004; Treichel et al., 2010

Lin et al. (1996) produced an alkaline lipase from Pseudomonas pseudoalcaligenes F-

111 in a medium that contained both olive oil (0.4%) and Triton X-100 (0.2%). The

addition of Triton X-100 enhanced the alkaline lipase production by 50-fold compared to

using olive oil alone. Kim et al. (1998) reported production of a highly alkaline

thermostable lipase by Bacillus stearothermophilus L1 in a medium that contained beef

tallow and palm oil. Salihu et al. (2011) reported the use of palm oil mill effluent for lipase

production by Candida cylindracea with an activity of 20.26 U/ml under the optimized

conditions.

Besides carbon sources, nitrogen sources have significant effect on lipase

production as it is directly related to the cell growth and division of microbial strains.

Nitrogen source can be provided in either inorganic (ammonium, sodium salts etc.) or

organic form (proteins, amino acids and urea etc.). Generally, organic nitrogen is preferred,

Microbial strain Incubation time

pH and Incubation temperature

References

Acinetobacter radioresistens - pH 7.0, 30 °C Chen et al. (1998)

Bacillus cereus 24 h pH 8.0, 30–33 °C Dutta and Ray (2009)

Bacillus cereus and B. coagulans 24 h - El-Shafei and Rezkallah (1997)

Bacillus clausii SKAL-16 - pH 8–10 Lee and Park (2008)

Bacillus coagulans BTS-3 - pH 8.5, 55 °C Kumar et al. (2005)

Bacillus sp. 36 h - Handelsman and Shoham (1994)

Bacillus sp. FH5 48 h pH 8.0, 37 °C Hasan et al. (2006)

Bacillus thermoleovorans IHI-91 - pH 6.0, 65 °C Markossian et al. (2000)

Candida rugosa 60 h - Song et al. (2001)

Cryptococcus sp. S-2. 120 h - Zhen-Jian and Chun-Yun (2009)

Lactobacillus plantarum DSMZ, 12028

- pH 5.5 Lopes et al. (1999)

Penicillium candidum - pH 9.0, 35 °C Ruiz et al. (2001)

Penicillium wortmanii 7 days - Costa and Peralta (1999); Song et al. (2001)

Pseudomonas sp. 7323 - pH 9.0, 30 °C Zhang and Zeng (2008)

Pseudomonas strain N 72 h - Sarkar et al. (1998)

Rhizopus oryzae 4 days - Kamini et al. (2000)

Salinivibrio sp. SA-2 - pH 8.0, 35 °C Amoozegar et al. (2008)

32

such as peptone and yeast extract, which have been used as nitrogen source for lipase

production by various Bacillus sp. such as Bacillus strain A30-1, Bacillus alcalophilus and

Bacillus licheniformis strain H1. Inorganic nitrogen sources such as ammonium chloride

and diammonium hydrogen phosphate have also been reported to be effective in some

microbes (Bradoo et al., 1999; Dong et al., 1999; Gilbert et al., 1991a, 1991b; Rathi et al.,

2001). Kempka et al. (2008) reported corn steep liquor, yeast extract and peptone as best

nitrogen sources for lipase production from Penicillium verrucosumin while Mahanta et al.

(2008) reported peptone, NH4Cl and NaNO3 in case of Pseudomonas aeruginosa PseA. The

carbon and nitrogen sources used for lipase production from different lipolytic

microorganisms are presented in table 2.4

Table: 2.4. Carbon and nitrogen sources used for lipase production from different

microorganisms

Microorganisms Carbon sources Nitrogen sources References

A. calcoaceticus Lactic acid, oleic acid

- Mahler et al. (2000)

A. calcoaceticus LP009 Tween-80

Tryptone, yeast extract

Pratuangdejkul and Dharmsthiti (2000)

Acinetobacter sp. Tween-80/ olive oil

- Barbaro et al. (2001)

Bacillus cepacia RGP-10

Glucose, mustard oil

NH4Cl, (NH4)2HPO4 Rathi et al. (2001)

Bacillus licheniformis B-42

Fructose, glucose, Ammonium molybdate

Bayoumi et al. (2007)

Bacillus alcalophilus Maltose, soybean meal Peptone, Yeast extract

Ghanem et al. (2000)

Bacillus licheniformis MTCC-10498

Cotton seed oil Peptone Sharma et al. (2012)

Bacillus licheniformis strain H1

Glucose

Peptone, yeast extract, beef extract

Khyami-Horani (1996)

Bacillus sp. Lactose Peptone Kumar et al. (2012)

Bacillus sp. Olive oil

Peptone, yeast extract

Sugihara et al. (1991)

Bacillus sp. Soybean flour, stearyl glycerol esters or natural fats

- Kambourova et al. (1996)

Bacillus sp. glucose, olive oil Yeast extract Heravi et al. (2008)

Bacillus sp. FH5 Salicin Yeast extract

Hasan et al. (2006)

Bacillus sp. RSJ1 Tween-80/ olive oil

Peptone, yeast Extract

Sharma et al. (2002b)

33

Sources: Modified from Gupta et al., 2004; Hasan et al., 2004; Treichel et al., 2010

2.4.2 Statistical optimization

While developing an industrial fermentation, designing the fermentation medium is

of critical importance because medium composition significantly affects product

concentration, yield and productivity. For commodity products, medium cost can

substantially affect the overall process economics. Designing the medium is a laborious,

expensive and often time-consuming process involving many experiments (Kennedy and

Krouse, 1999). There is a general practice of determining optimal concentration of media

components by varying one factor at a time. However, this method does not depict the net

effect of total interactions among the various media components (Rathi et al., 2001). Thus,

the emphasis has shifted towards medium optimization using statistical methods.

The factorial design of a limited set of variables is advantageous in relation to the

conventional method of manipulation of a single parameter per trial, as the latter approach

frequently fails to locate the optimal conditions for the process, due to its failure to

Bacillus sp. strain 398

Glycerol

Polypeptone, yeast extract, beef extract

Kim et al. (1994)

Bacillus sp., Pseudomonas sp.

Dextrose, Triolein

Tryptone, yeast Extract

Lanser et al. (2002)

Bacillus strain A30-1 (ATCC 53841)

Corn oil

Ammonium chloride, yeast extract

Wang et al. (1995)

Bacillus THL027 (thermophilic)

glucose, rice bran oil, rice bran

- Dharmsthiti and Luchai (1999)

Candida cylindracea Tween-80 , Palm oil Peptone Salihu el al. (2011)

Candida rugosa glucose, olive oil - Song et al. (2001)

Pseudomonas aeruginosa EF2

Tween-80

KNO3 Gilbert et al. (1991b)

Pseudomonas aeruginosa LP602

Whey, soybean oil, Glucose

Ammonium sulphate, yeast extract

Dharmsthiti and Kuhasuntisuk (1998)

Pseudomonas sp. Ground soybean, soluble starch

Corn steep liquor, NaNO3

Dong et al. (1999)

Pseudomonas sp. Soya peptone, Cottonseed meal

Soya peptone

Kulkarni and Gadre (1999)

Pseudomonas sp. G6 n-Hexadecane, tributyrin n.s.

Kanwar et al. (2002)

Pseudomonas sp. strain KB 700A (recombinant lipase)

Casamino acids

Yeast extract

Rashid et al. (2001)

Serratia marcescens Sucrose, glycine Peptone, K2HPO4 Su et al. (2011)

34

consider the effect of possible interactions between factors. Moreover, the factorial design

makes it possible to take advantage of practical knowledge about the process during the

final RSM analysis (Kalil et al., 2000). An efficient and widely used approach is the

application of Plackett–Burman (PB) designs that allow efficient screening of key variables

for further optimization in a rational way.

An alkaline lipase from Burkholderia multivorans was produced after 15 h of

cultivation in a 14-L bioreactor. The medium optimization led to an increase of 12-fold in

lipase production. Initially, the effect of nine factors i.e. concentrations of glucose, dextran,

olive oil, NH4Cl, trace metals, K2HPO4, MgCl2, and CaCl2 and inoculum density were studied

using the PB experimental design. After screening of the most significant factors by the PB

design, optimization was carried out in terms of the concentration of olive oil, glucose, and

yeast extract, inoculum density, and fermentation time. The optimal medium composition

for the lipase production was determined to be (% w/v): glucose 0.1, olive oil 3.0, NH4Cl

0.5, yeast extract 0.36, K2HPO4 0.1, MgCl2 0.01, and CaCl2 0.4 mM (Gupta et al., 2007).

Dandavate et al. (2012) statistically optimized the medium components and

enhanced the lipase production by 2.2 fold from bacterium Burkholderia multivorans V2.

Salihu el al. (2011) used the sequential optimization strategy based on statistical

experimental design including one-factor-at-a-time method to enhance the production of

lipase by Candida cylindracea ATCC 14830 using palm oil mill effluent as a basal medium in

shake flask cultures. They employed the two-level Plackett–Burman (PB) design to screen

the medium components that significantly influenced the production. Following the one-

factor-at-a-time method, they identified three significant components influencing lipase

production as peptone, Tween-80 and inoculum followed by RSM based on the face-

centered central composite design to find the optimum values of these three components

i.e 0.45 % (w/v) peptone, 0.65 % (v/v) Tween-80 and 2.2 % (v/v) inoculum. This optimum

medium led to a maximum lipase production of 20.26 U/ml, which was 5.19-fold higher

than the unoptimized medium.

Hasan-Beikdashti et al. (2012) optimized the production of an extracellular lipase

from the newly isolated bacterium Stenotrophomonas maltophilia via the statistical design

method. The initial screening of 10 factors of the medium components were done using

Plackett– Burman design, to find out the more significant factors viz. olive oil, peptone,

35

yeast extract, and ferrous sulfate. The level of each factor was subjected to optimization by

using the Box–Behnken technique, and a 9.1-fold enhancement of lipase productivity (from

500 U/ml to 4559 U/ml) was achieved overall in the presence of optimum levels of the

effective factors.

Kumar and Gupta (2008) compared the medium optimization for the yeast T. asahii

by both one variable at a time and statistical approach. A Plackett–Burman design for seven

independent variables (glucose, olive oil, yeast extract, malt extract, MgCl2, and CaCl2

concentrations and time) was applied to select the most significant factor. Wang et al.

(2008) optimized the fermentation medium for lipase production by Rhizopus chinensis. In

order to improve the productivity of lipase, the effects of oils and oil-related substrates

were assessed by orthogonal test and response surface methodology (RSM). The optimized

medium for improved lipase activity consisted of peptone, olive oil, maltose, K2HPO4, and

MgSO4.7H2O. Rajendran et al. (2008) used the Plackett–Burman statistical experimental

design to evaluate the fermentation medium components. The effect of 12 medium

components was studied in 16 experimental trials. Glucose, olive oil and peptone were

found to have more significant influence on lipase production by Candida rugosa.

Ruchi et al. (2008) carried out media optimization through RSM for cost-effective

production of lipase by Pseudomonas aeruginosa. The effects of 11 media components

(peptone, tryptone, NH4Cl, NaNO3, yeast extract, glucose, glycerol, xylose, arabic gum,

MgSO4, and NaCl) were assessed by a Plackett–Burman design, and the most significant

factors (arabic gum, MgSO4, tryptone, and yeast extract) optimized by the RSM. After

optimization, the lipase production was increased 5.58-fold, yielding an activity of 4,580 U

mL−1. Kaushik et al. (2006) used the RSM approach to investigate the production of an

extracellular lipase from Aspergillus carneus. Interactions were evaluated for five different

variables (sunflower oil, glucose, peptone, agitation rate, and incubation period) and 1.8-

fold increase in production was reported under optimized conditions.

He and Tan (2006) used RSM to optimize the culture medium for lipase production

by Candida sp. 99-125. Firstly, a Plackett– Burman design was used to evaluate the effects

of different components of the culture medium (soybean oil, soybean meal, K2HPO4,

KH2PO4, (NH4)2SO4, MgSO4, and Spam 60). Soybean oil, soybean meal, and K2HPO4

concentrations significantly influenced the lipase production. Results were optimized using

36

central composite designs and response surface analysis. The optimized conditions

resulted in an increase in lipase production from 5,000 to 6,230 U mL−1 in a shaken flask

system. Burkert et al. (2004) studied the effect of carbon source (soybean oil, olive oil, and

glucose) and nitrogen source (corn steep liquor and NH4NO3) on lipase production by

Geotrichum sp. using RSM reaching a lipase activity of 20 U mL−1.

2.5 Purification of lipase

Many lipases have been extensively purified and characterized in terms of their

activity and stability profiles. The purification of lipase from different microorganisms has

been reported through several techniques such as precipitation, hydrophobic interaction

chromatography, gel filtration, ion exchange chromatography and affinity chromatography

as shown in Table 2.5.

Tamilarasana et al. (2012) purified extracellular lipase from Bacillus sphaericus

MTCC by DEAE–Sepharose anion exchange chromatography resulting in 377 U/mg specific

activity and 17.33-fold purification with 5.7 % recovery. The molecular weight of the

purified lipase was determined to be 69 kDa using SDS-PAGE.

Cao et al. (2012) have purified organic solvent-stable lipase from newly isolated

solvent-tolerant bacterium Pseudomonas stutzeri LC28 by acetone precipitation and anion

exchange chromatography. The apparent molecular mass of the purified lipase was 32 kDa

as estimated by SDS-PAGE.

Velu et al. (2012) purified a thermostable lipase produced from the bacterium

Aeromonas caviae AU04 by 3.3-fold with 28.7% recovery using ammonium sulphate

precipitation and hydrophobic interaction chromatography.

Romero et al. (2012) purified an extracellular lipase from Staphylococcus aureus

using PALL’S Microsep centrifugal device (10 kD cut off), hydrophobic interaction (phenyl

sepharose CL-4B column) and Superose-12 gel filtration chromatography and found to

have a molecular mass of ~ 49 kDa.

Masomian et al. (2012) purified a thermostable and organic solvent-tolerant lipase

(MW 50 kDa) produced by Aneurinibacillus thermoaerophilus strain HZ using anion

exchange chromatography on Q-Sepharose and gel filtration on Sephadex-G75. A final

37

specific activity of 43.5 U/mg was obtained with an overall recovery of 19.7 % and 15.6-

fold purification.

Shaoxin et al. (2007) purified lipase from Bacillus cereus C71 to homogeneity by

ammonium sulfate precipitation, followed by Phenyl-Sepharose chromatography, DEAE

ion-exchange chromatography and CIM QA chromatography. This purification procedure

resulted in 1092-fold purification of lipase with 18 % yield. The molecular mass of the

purified enzyme was found to be ~42 kDa by SDS-PAGE.

Yu et al. (2007) purified an extracellular lipase from Yarrowia lipolytica (Lip2) by

ion-exchange chromatography on Q- Sepharose FF, followed by hydrophobic interaction

chromatography on Butyl-Sepharose FF. SDS-PAGE showed its molecular weight as 38 kDa.

Shu et al. (2006) purified an extracellular lipase from Antrodia cinnamomea BCRC

35396 by ammonium sulphate precipitation and DEAE-Sepharose chromatography. The

yield and purified factor were 33.7 % and 17.2- fold, respectively.

Kumar et al. (2005) purified lipase from thermophilic and alkaliphilic Bacillus

coagulans BTS-3 to homogeneity by 40-fold using ammonium sulfate precipitation and

DEAE–Sepharose column chromatography. Its molecular weight was 31 kDa on SDS–PAGE.

Lianghua et al. (2005) reported the purification of an extracellular lipase from the

fermentation broth of Bacillus coagulans ZJU318 by CM-Sepharose chromatography

followed by Sephacryl S-200 chromatography. The lipase was purified 14.7-fold with 18 %

recovery and a specific activity of 141.1 U/mg. The molecular weight of the homogeneous

enzyme was 32 kDa as determined by SDS-PAGE.

The lipase from Pseudomonas fluorescens HU380 was purified by Phenyl-Toyopearl

fractionation, DEAE-Sepharose chromatography and Superdex-200HR chromatography

(Kojima et al., 2004). The enzyme was purified 24.3-fold with 14 % yield and 9854 U/mg

specific activity having a molecular weight of 64 kDa.

Hiol et al. (2000) purified an extracellular lipase produced by Rhizopus oryzae using

ammonium sulfate precipitation, Sulfopropyl-Sepharose chromatography, Sephadex G-75

gel filtration, and a second Sulfopropyl-Sepharose chromatography step. The enzyme was

purified 1200-fold and had a molecular mass of 32 kDa determined by SDS-PAGE and gel

filtration.

38

A thermostable lipase produced by a thermophilic Bacillus sp. J 33 was purified to

175-fold by ammonium sulfate and Phenyl-Sepharose column chromatography (Nawani

and Kaur, 2000) with a recovery of 15.6 %. The enzyme was shown to be a monomeric

protein of 45 kDa. The enzyme hydrolyzed triolein at all the positions.

A three-step procedure involving ammonium sulfate precipitation, DEAE- Sephacel

ion exchange chromatography, and Sephacryl S-200 gel filtration chromatography was

used to purify a lipase from Bacillus thermoleovorans ID-1 to homogeneity (Lee et al.,

1999). The protein was purified 223-fold with molecular mass of 34 kDa as determined by

SDS-PAGE.

An extracellular lipase from Pseudomonas aeruginosa KKA-5 was purified using

ammonium sulfate precipitation and successive chromatographic separations on

hydroxyapatite (Sharon et al., 1998). After 5l8-fold purification, the enzyme was

homogenous electrophoretically and its molecular mass was estimated to be 30 kDa.

Kim et al. (1996) purified a highly alkaline extracellular lipase of Proteus vulgaris by

ion-exchange chromatography. The purified lipase had a maximum hydrolytic activity at

pH 10.0 and its molecular mass was 31 kDa as determined by SDS-PAGE.

Lin et al. (1996) purified an alkaline lipase from Pseudomonas pseudoalcaligenes F -

111 to homogeneity. Its apparent molecular mass by SDS-PAGE was 32 kDa and the

isoelectric pH was 7.3.

Chartrain et al. (1993) purified a lipase from Pseudomonas aeruginosa MB5001

using a three-step procedure i.e. concentration by ultrafiltration followed by ion-exchange

chromatography and gel filtration. The purified lipase had a molecular mass of 29 kDa and

exhibited maximum activity at 55°C and pH 8.0.

2.5.1 Characterization of purified lipase

Purified lipases have been characterized for various physico-chemical properties.

Primary structures of several lipases have been determined either from amino acid or

nucleic acid sequences. Lipases sequenced to date share sequence homologies including

conserved region Gly-X-Ser-X-Gly. The serine residue is suspected to be essential for

binding to lipid substrates (Antonian, 1988). The molecular weight and pH and

39

temperature stability of some purified lipases from different microorganisms are listed in

Table 2.5

2.5.1.1 Molecular weight

The molecular weights of lipases from different organisms are shown in Table 2.5.

Lipases have been found to be glycoproteins differing in their molecular weight. In case of

Bacillus sp., the molecular weight may be as low as 11.6 kDa in B. stearothermophilus (Kim

et al., 20000 and as high as 75 kDa in B. pumilus (Jose and Kurup, 1999). Similarly, lipases

purified from other microorganisms exhibited a lot of variation with regard to their

molecular weights as seen from Table 2.5.

2.5.1.2 Effect of pH and temperature

Generally, bacterial lipases have neutral (Dharmsthiti et al., 1998; Dharmsthiti and

Luchai, 1999; Lee et al., 1999) or alkaline pH optima (Kanwar and Goswami, 2002;

Schmidt-Dannert et al., 1994; Sidhu et al., 1998a, 1998b; Sunna et al., 2002) with the

exception of Pseudomonas fluorescens SIK W1 lipase, which exhibited an acidic optimum at

pH 4.8 (Andersson et al., 1979). Lipases from Bacillus stearothermophilus SB-1, Bacillus

atrophaeus SB-2 and Bacillus licheniformis SB-3 were active over a broad pH range (pH 3.0–

12.0; Bradoo et al., 1999). Bacterial lipases possess stability over a wide range, from pH 4.0

to 11.0 (Dong et al., 1999; Kojima et al., 1994; Wang et al., 1995).

The rate of enzyme catalyzed reactions approximately doubles for each 10 °C

increase in temperature. Assuming the enzyme is stable at elevated temperatures, the

productivity of the reaction can be enhanced greatly by operating at a relatively high

temperature but most of the lipases become deactivated beyond 45°C. Thus thermal

stability is a desirable characteristic of lipases (Janssen et al., 1994).

Thermostable lipases have been isolated from many sources, including

Pseudomonas fluorescens (Kojima et al., 1994); Bacillus sp. (Wang et al., 1995; Sidhu et al.,

1998a, b), Bacillus coagulans and Bacillus cereus (El-Shafei and Rezkallah, 1997);

Geotrichum sp. and Aeromonas sobria (Lotrakul and Dhannsthiti, 1997; Macedo et al.,

1997). Many researchers reported highly thermostable lipases (Gao and Breuil, 1995; Kim

et al., 1998; Lee et al., 1999). It was suggested that thermal stability of a lipase was related

40

with its structure (Zhu et al., 2001). Thermostability is influenced by environmental factors

such as pH and the presence of metal ions. At least in some cases, thermal denaturation

appeared to occur through intermediate states of unfolding of the polypeptide (Zhu et al.,

2001). Mutations in the ‘lid’ region of the enzyme could significantly affect heat stability.

Attempts are being made to engineer lipase protein for improved thermal stability.

Compared to the native enzyme, thermal and operational stability of many lipases could be

significantly enhanced by immobilization (Arroyo et al., 1999; Hiol et aI., 2000; Xu et al.,

1995). Candida antarctica lipase B could be thermally stabilized by immobilization. The

native enzyme and the covalently immobilized preparation appeared to follow different

modes of thermal deactivation (Arroyo et al., 1999).

2.5.1.3 Effect of metal ions and other compounds on lipase activity

There are many reports related to the effect of various metal ions on lipase activity.

However, the concentration of metal ion, extent and mechanism of induction could vary

from species to species (Saxena et al., 1994) and in some case, same metal ion might also

act as suppressor. The purified lipase of Bacillus coagulans BTS-3 was found to be inhibited

by Al3+, Co2+, Mn2+, and Zn2+ ions while K+, Fe3+, Hg2+ and Mg2+ ions enhanced the enzyme

activity; Na+ ions had no effect on enzyme activity (Kumar et al., 2005). The lipase of an

Alaskan psychrotroph Pseudomonas sp. B11-1 was strongly inhibited by Zn2+, Cu2+, Fe3+,

and Hg2+ but was not affected by phenylmethylsulfonyl fluoride (PMSF) and bis-

nitrophenyl phosphate. Tamilarasana et al. (2012) reported the enhancement in lipase

activity in the presence of Mg2+, Ca2+, Cu2+, K2+ and Tween 20. Chartrain et al. (1993)

observed that an extracellular lipase of Pseudomonas aeruginosa MB5001 was strongly

inhibited by 1 mM ZnS04 (94 % inhibition) but was stimulated by adding 10 mM CaCl2

(1.24-fold stimulation) and 200 mM taurocholic acid (1.6-fold stimulation). The lipase

activity was not affected by Ca2+, Mg2+, Mn2+, Na+, K+, Cu2+, EDTA, p-chloromercuribenzoic

acid, and iodoacetate while the enzyme was inhibited by Ag+, Fe2+, Hg2+, and isopropyl

fluorophosphates (Mase et al., 1995). In another similar study with metal ions (1 mM) and

chelating agents, Pseudomonas pseudoalcaligenes F –III lipase activity was 60 % inhibited

by Fe3+ but not by Ca2+, Hg2+, Zn2+, Mn2+, Cu2+, Mg2+, Co2+, Cd2+, and Pb2+ (Lin et al., 1996).

41

Microorganisms MW (kDa)

pH and temperature stability

Purification steps Recovery (%)

Purification fold

References

Acinetobacter calcoaceticus

30.5 30– 40°C - - Brune and Gotz, (1992)

Acinetobacter sp. RAG-1

33 Active at temperatures up to 70 °C

- - - Snellman et al. (2002)

Aeromonas caviae AU04 - - Ammonium sulphate precipi-tation and Hydrophobic interaction chromatography

28.7 3.3 Velu et al. (2012)

Aneurinibacillus thermoaerophilus strain HZ

50 Broad pH stability (4-9), 30-55 °C

Anion exchange chromatography on Q-Sepharose and gel filtration on Sephadex-G75

19.7 15.6 Masomian et al. (2012)

Bacillus licheniformis strain H1

-

Stable at alkaline pH 9–11, 65% residual activity at pH 12 after 30 min at 4°C, retained 100% activity after 15 min at 70°C

- - - Khyami-Horani (1996)

Bacillus alcalophilus

- Stable at pH 10.0–10.5, 80% activity at pH 11.0 after 1 h; stable at 60°C for 1 h, 70% residual activity at 75°C

Ammonium sulfate precipitation and Sephadex G-100

- 111 Ghanem et al. (2000)

Bacillus licheniformis MTCC 6824 w

- Ammonium sulphate precipi-tation, ether precipitation, dialysis followed anion exchange chromatography on amberlite IRA 410 A and gel exclusion on Sephadex G 100

8.36 208 Chakrabort et al. 2008

Bacillus pumilus 75 - Ammonium sulfate fractionation and gel filtration on Sephadex

- 100

Jose and Kurup (1999)

Table 2.5 Purification and characterization of various lipases produced by microorganisms

42

Bacillus pumilus RK31 62 Gel filtration chromatography using Sephadex G200 and Ion exchange chromatography with DEAE Cellulose

186 Kumar et al. 2012

Bacillus sp. 22 Stable over pH 5.0–11.5, stable at 65°C for 30 min at pH 5.6

Ammonium sulfate fractionation, treatment with acrinol, DEAE- Sephadex A-50, Toyopearl HW-55F and butyl-Toyopearl 650 M

9 7762 Sugihara et al. (1991)

Bacillus sp. 25 - Acetone fractionation, two acetone precipitations and Octyl-Sepharose CL-4B, Q-Sepharose and Sepharose-12

20 3028 Imamura and Kitaura (2000)

Bacillus sp. THL 027 69 Stable over pH 6.0−8.0, 80% residual activity after 1 h at 75°C

Ultrafiltration and Sephadex G-100

- 2.6 Dharmsthiti and Luchai (1999)

Bacillus sphaericus 205y

30 Broad pH stability (5-13), 37-55 °C

Ultrafiltration and hydrophobic interaction chromatography (HIC)

32

8 Sulog et al. (2006)

Bacillus stearo-thermophilus

11.6 - CM-Sepharose and DEAE-Sepharose

62.2 Kim et al. (2000)

Bacillus strain A30-1

65 90–95% residual activity after 15 h at pH 5.0–10.5, half-life of 8 h at 75°C

- - - Wang et al. (1995)

Bacillus subtilis 168 19 Stable at pH 12; 100 % activity after 30 min. at 40°C

- - - Lesuisse et al. (1993)

Bacillus thermocatenulates

16 - Disintegration, heat precipitation, ion-exchange and hydrophobic interaction chromatography

- 312 Schmidt-Dannert et al. (1996)

Pseudomonas aeruginosa

30 - Ammonium sulfate precipitation, hydroxyapatite column chromatography

518 Sharon et al. (1998)

43

Pseudomonas cepacia 60 - Polyoxyethylene detergent 14EO6- based aqueous two-phase partitioning

76 24

Terstappen et al. (1992)

Pseudomonas cepacia l

58 - Liquid–liquid (10% PEG 6000 and 10% Dextran 500) extraction and chromatography using Q-Sepharose

30 55 Dunhaupt et al. (1991)

Pseudomonas fluorescens

45 - Ammonium sulfate precipitation and chromatography on DEAE-Cellulose and Octyl-Sepharose CL-4B

21 3390 Sztajer et al. (1992)

Pseudomonas fluorescens

33 - Ultrafiltration, ammonium sulfate precipitation, DEAE-Toyopearl 650 M and Phenyl-Toyopearl 650

42 6.1

Kojima et al. (1994)

Pseudomonas fragi 33 - Acidification, ammonium sulfate fractionation, DEAE-Toyopearl 650M and DEAE-Sepharose CL-6B

48 68 Nishio et al. (1987)

Pseudomonas putida 3SK

45 - DEAE-Sephadex A-50 and Sephadex G-100

21 5.3

Lee and Rhee (1993)

Pseudomonas spp. ATCC 21808

35 - Q-Sepharose, Octyl-Sepharose and the enzyme eluted with isopropanol

56 159

Kordel et al. (1991)

Rhizopus arrhizus 67 - Ammonium sulfate fractionation and Sephadex G-100

42 720

Chattopadhyay et al. (1999)

Rhizopus chinensis 28.4 - Ether, Toyopearl 650M, Super Q-Toyopearl and CM-Cellulofine C-500

27.6

Yasuda et al. (2000)

Rhizopus delemar 30.3 - Affinity chromatography and CM-Sephadex

30 10.3

Haas et al. (1992)

44

Rhizopus japonicas NR 400

30 - Hydroxyapatite, Octyl-Sepharose and Sephacryl S-200

31 93

Suzuki et al. (1986)

Rhizopus oryzae - Acetone precipitation (80%), Sephadex G-100

64 160

Razak et al. (1997)

Rhizopus oryzae 32 - Ammonium sulfate fractionation, Sulfopropyl-Sepharose, Sephadex G-75 and again on Sulfopropyl-Sepharose

22 1260

Hiol et al. (2000)

Staphylococcus aureus 49 - PALL’S Microsep centrifugal device (10 kDa cut off), phenyl sepharose CL-4B column, Superose-12 gel filtration chromatography

- - Romero et al. (2012)

45

Hiol et al. (2000) studied the effect of various compounds and enzyme inhibitors on

Rhizopus oryzae lipase. Among the metal ions, Fe2+, Fe3+, Hg2+, and Cu2+ ions strongly

inhibited the enzyme but benzamidine and PMSF had no effect on the enzyme activity.

2.5.1.4 Enzyme kinetics parameters (Km and Vmax)

Michaelis-Menten constant i.e. Km is specific for a specific enzyme and denotes the

enzyme specificity or affinity towards the substrate. The Km value depends on the type of

substrate used for enzyme assay. The unit of Km representation for lipase is mg/mL but no

uniformity has been observed in the unit for Vmax representation which makes their

comparison complicated. The value of Km, if known, can be used to improve the assay

conditions by changing the substrate concentration so that it should not be limiting. The

lipase purified from Pseudomonas fluorescens strain AFT 36 had a Km of 3.65 mM with

tributyrin as substrate and was inhibited by concentrations of substrate greater than

approximately 17 mM (Fox and Stepaniak, 1983). The Km of lipase from Brazilian strain of

Fusarium solani FSI using p-NPP (p-nitrophenyl palmitate) as substrate was 1.8 mM with a

Vmax of 1.7 μmol/min/mg protein (Maia et al., 2001). The Km and Vmax values of the lipase

purified from Bacillus sp. FH5 were 5.05 mM and 0.416 μmol/mL/min, respectively (Hasan

et al., 2006). The Km of the lipase purified from Serratia marcescens was 1.35 mM on

tributyrin (Abdou, 2003). The values of Km and Vmax of the lipase from Aspergillus niger

F044 using p-NPP as substrate were 7.37 mM and 25.91 μmol/min/mg, respectively (Shu

et al., 2007).

2.6 Immobilization of lipase

Immobilization means restricting the mobility of biocatalysts either completely or to

a small limited region through various means such as attachment to an insoluble matrix by

adsorption, ionic and covalent forces, cross-linking, adsorption followed by cross-linking,

physical entrapment, microencapsulation etc. For practical applications, immobilization of

enzymes on solid materials may offer several advantages, which include reusability of

enzyme, ease of product separation, improvement of enzyme stability, continuous use of

enzyme in a continuous reactor system, reduced effluent disposal problem, and

development of multienzyme reaction systems. Immobilization often stabilizes structure of

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the enzymes, thereby allowing their applications even under harsh environmental

conditions of pH, temperature and organic solvents, and thus enables their uses at high

temperatures, in non-aqueous media, and in the development of continuous processes

allowing more economic organization of the operations. The major contribution to achieve

a good performance of immobilized catalyst is primarily provided by the strategy employed

for immobilization (Cardias et al., 1999) and by the characteristics of the support.

The desirable characteristics of solid supports used for immobilization include large

surface area, low cost, reusability, good chemical, mechanical and thermal stability and

insolubility (Miletic et al., 2012).

Lipases have been immobilized by different processes, such as physical adsorption,

covalent attachment, entrapment and microencapsulation using various supports

(Bhushan et al., 2008; Brem et al., 2012; Krajewska et.al., 2004; Mateo et al., 2007; Pavlidis

et al., 2010; Saunders and Brask, 2010; Zhao et al., 2009).

Covalent immobilization involves multipoint covalent protein attachment directly or

through a spacer to a carrier. One of the benefits of the method is that enzyme leaching is not

possible. Lee et al. (2009) have suggested the use of nano-sized magnetite particles for

immobilizing lipase. Carbon nano tubes have found extensive applications as biosensors and

nano-biocatalysts owing to their application as immobilization support for enzymes (Lee et al.,

2010). Electrospun nanofibres proved to be excellent immobilization supports and lipase was

immobilized by physical adsorption on to electrospun nanofibres (Sakai et al., 2010) and

magnetic silica nanocomposite particles (Kalantari et al., 2012; Kuwahara et al., 2012).

Li et al. (2013) adsorbed Yarrowia lipolytica lipase onto the functionalized

multiwalled carbon nanotubes (MWNTs) i.e amino-cyclodextrin was covalently attached to

MWNTS. The immobilized lipase was utilized for the resolution of the model compound (R,

S)-1-phenyl ethanol in heptane, the ionic liquid [Bmim]PF6 i.e (1-Butyl-3-

methylimidazolium hexafluorophosphate) as well as the heptane/[Bmim]PF6 mixture. In

the reaction media, the enzymatic activity of the immobilized lipase was much higher than

that of the native lipase. In comparison to the catalysis in the ionic liquid and heptane,

when using the mixture of heptane/[Bmim]PF6 as the reaction medium, the catalysis by

the immobilized lipase at the heptane–ionic liquid interface exhibited a higher catalysis

activity.

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Kima et al. (2013) immobilized Staphylococcus haemolyticus L62 lipase on a poly

(methacrylate-co-divinyl benzene) resin by two different methods: hydrophobic

adsorption/entrapment (H-L62) and hydrophobic adsorption/entrapment plus covalent cross-

linking (HC-L62). Both the immobilized enzymes showed quite increased temperature and pH

stabilities, whereas they had very similar optimal temperature and pH in comparison with the

soluble free enzyme.

Zheng et al. (2012) immobilized Candida rugosa lipase (CRL) on the hydrophobic

magnetic microspheres via the active epoxy groups. The resulting immobilized CRL had

better resistance to pH and temperature inactivation in comparison to free CRL, the

adaptive pH and temperature ranges of lipase were widened, and it exhibited good thermal

stability and reusability. The immobilized CRL was used as biocatalyst for enzymatic

esterification of phytosterols with unsaturated fatty acids (UFAs) to produce the

corresponding phytosterol esters, which were also converted in high yields to the

corresponding long-chain acyl esters via transesterification with methyl esters of fatty

acids or triacylglycerols using magnetic immobilized CRL as biocatalyst.

Brem et al. (2012) immobilized lipase AK “Amano” 20 from Pseudomonas fluorescens

using diverse immobilization techniques such as sol–gel entrapment, cross-linked enzyme

aggregates, adsorption etc. These immobilization methods enabled the fine tuning of enzymatic

activity and enantioselectivity for the kinetic resolution of racemic ethyl 3-aryl-3-

hydroxypropanoates. The immobilized lipase was efficiently reused for ten cycles with high

enantioselectivity.

Bhushan et al. (2007) immobilised Arthrobacter sp. (RRLJ-1, named ABL) on a series

of synthetic macroporous epoxy copolymers beads with varying pore sizes, surface area

and hydrophobicity. The copolymer poly(glycidyl methacrylate-co-ethylene

dimethacrylate) beads, with 75 % crosslink density and 10 % of epoxy groups modified

with dibutyl amine had a pore volume of 0.77 mL/g. The immobilized ABL displayed

enhanced thermal, organic solvent and pH stability compared to the free enzyme. The

immobilized enzyme was used repeatedly (15 cycles) to resolve the racemic fluoxitine

intermediate (ethyl-3-hydroxy-3-phenyl propanoate) without any loss in stereospecificity.

Chiou et al. (2004) reported a method for covalent attachment of Candida rugosa lipase

to two types of chitosan beads by activating the hydroxyl groups using carbodiimide as the

48

coupling agent. Immobilization enhanced the enzyme stability against changes of pH and

temperature and increased enzyme activity up to 110 %. Immobilized lipase using dry and wet

chitosan beads retained 78 % and 85 % of its initial activity after ten cycles.

Jaeger and Reetz (1998) produced glutaraldehyde cross-linked microcrystals of Candida

rugosa lipase. These cross-linked crystals were used for the chiral resolution of commercially

important compounds by ester hydrolysis.

Kuo et al. (2012) used Fe3O4–chitosan nanoparticles for the covalent immobilization of

lipase from Candida rugosa using N-(3-dimethylaminopropyl)-N-ethylcarbodiimide and N-

hydroxysuccinimide as coupling agents. Response surface methodology was employed to search

the optimal immobilization conditions and understand the significance of the factors affecting the

immobilized lipase activity. Based on the ridge max analysis, the optimum immobilization

conditions were immobilization time 2.14 h, pH 6.37, and enzyme/support ratio 0.73 (w/w); the

highest activity obtained was 20 U/g Fe3O4–chitosan. After twenty repeated uses, the

immobilized lipase retained over 83 % of its original activity. The immobilized lipase showed

better operational stability, including wider thermal and pH ranges, and remained stable after 13

days of storage at 25 °C.

Ondul et al. (2012) immobilized Candida antarctica A and Thermomyces lanuginosus

lipases on cotton terry cloth fibrils using polyethyleneimine. The highest amount of enzyme

precipitate was obtained at the 0.2 % PEI to enzyme ratio of 1/20–1/40 for both lipases. At pH

values below 8, aggregation and precipitation did not occur for C. antarctica A lipase whereas

pH did not affect PEI–enzyme aggregate formation for T. lanuginosus lipase. Immobilized

enzyme amount was approximately 180 mg/g support and 200mg/g support for T. lanuginosus

and C. antarctica A lipase, respectively. Immobilization had no effect on the optimum

temperature and it was 60 ◦C for both free and immobilized enzymes. Immobilized lipases

exhibited better operational and storage stability and could be stored at room temperature with a

little activity lost during 28 days.

Yucel (2012) immobilized lipase from Thermomyces lanuginosus onto olive pomace.

Response surface methodology was used to optimize the conditions for the maximum

activity and to understand the significance and interaction of the factors affecting the

specific activity. 5-level-3-factor central composite design was employed to evaluate the

effects of immobilization parameters such as enzyme concentration (3–15 %, v/v), pH

49

(5.0–9.0) and buffer concentration (20–100 mM) on the specific activity of immobilized

lipase. The predicted specific activity was 6.0 mmol p-NP/mg enzyme min under the

optimal conditions. Immobilized lipase was stable retaining >80 % activity after being used

repeatedly for 10 consecutive batches of pomace oil transesterification.

Bhushan et al. (2008) immobilized Arthrobacter sp. lipase (ABL) on various synthetic

macroporous alkylated glycidyl epoxy copolymers with varying hydrophobicity, pore volume

and surface area. Among all the polymers prepared and used only two epoxy polymers GMA-

EGDM 75-20(I) and GMA-EGDM 75-30(I) with particle size in the range of 150–450 nm,

epoxy groups 80 and 70 %, tertiary amino groups 20 and 30% was found suitable for

immobilization. The immobilized enzyme matrices were tested for the hydrolysis of triglycerides

using tributyrin as substrate as well as for racemic resolution of ethyl-3-hydroxy-3-phenyl

propanoate (fluoxetine intermediate, an antidepressant drug) and racemic chiral auxiliary, acetyl-

1-phenyl ethanol (intermediate of many chiral drugs). These immobilized lipase matrices were

recycled (15 cycles) with very high stability on recycling, high-enantioselectivity, high

conversion and faster recovery of product as compared to free enzyme.

2.7 Biotechnological Applications of Lipases

The industrial applications of lipases have been reviewed by many researchers

(Adriano et al., 2012; Hasan et al., 2006; Horchani et al., 2012; Kapoor et al., 2012; Thakur

et al., 2012; Yu et al., 2012). Lipases can be used as biocatalyst in a variety of applications such

as resolution of drugs, esterification/transesterification reactions etc. (Iso et al., 2001). The

biosynthesis of esters is currently of much commercial interest because of the increasing

popularity and demand for natural products amongst consumer. Biotransformations and

enzymatic methods of ester synthesis are more effective when performed in non-aqueous media

(Chand et al., 1997). Some of the applications of lipase are summarized below: -

2.7.1 Food industry

Lipases are employed in situ, and sometimes together with other enzymes, during

the elaboration of bread, cheese, and other foods to improve their shelf-life and their

rheological properties, or to produce aromas. Moreover, they are used ex situ to produce

flavours, and to modify the structure or composition of acylglycerols by inter- or

50

transesterification, in order to obtain acylglycerols with an increased nutritional value, or

suitable for parenteral feeding (Hita et al., 2009; Kim et al., 2010; Nadia et al., 2010).

2.7.2 Organic chemistry

Organic chemistry is the most important application of lipases after the food

industry. They are used to produce specific products that cannot be produced chemically,

or whose elaboration by classical chemical means is difficult or expensive. For example,

they are used in pharmaceutical and agrochemical industries for the modification or

synthesis of antibiotics, anti-inflammatory compounds, pesticides, etc., and for the

production of enantiopure compounds or the resolution of racemic mixtures (Hasan et al.,

2006; Li et al., 2011a; Lin et al., 2011; Pandey et al, 1999; Reetz, 2002).

2.7.3 Chiral resolution

Chirality is a geometrical attribute. An object that is not superimposable on its

mirror image is said to be chiral. The most common type of chiral organic molecule

contains a tetrahedral carbon atom attached to four different groups. Such a carbon is said

to be a stereogenic center and such a molecule exists in two stereoisomeric forms. Chirality

is not a prerequisite for bioactivity but in bioactive molecules where a stereogenic center is

present, great differences are observed in the activity of the enantiomers. This is general

phenomenon and applies to all bioactive substances, such as drugs, insecticides, herbicides,

flavors, fragrances and food additives. Conventional chemical synthesis of drugs containing

a chiral centre generally yields equal mixtures of enantiomers. During the past decade,

many studies have shown that racemic drugs usually have the desired therapeutic activity

residing mainly in one of the enantiomers and the other enantiomers might interact with

different receptor sites, which can cause unwanted side effects. The lipases have been

reported to accept a wide range of substrates for the production of compounds in high

enantiomeric excess which could be used as chiral building blocks for the synthesis of

compounds of pharmaceutical interest (Anand et al., 2004; Chaubey et al., 2006; Indu et al.,

2008; Indu et al. 2011; Kapoor et al., 2003; Padmapriya et al., 2011).

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2.7.4 Lipase as biosensor

The quantitative determination of triacylglycerol is of great importance in clinical

diagnosis and in food industry. The lipid sensing device as a biosensor is rather cheaper

and less time consuming as compared to the chemical methods for the determination of

triacylglycerols. An analytical biosensor was developed for the determination of lipids for

the clinical diagnosis (Masahiko et al., 1995). C. rugusa lipase biosensor from Candida

rugosa has been developed as a DNA probe (Benjamin and Pandey, 2001).

2.7.5 Lipases in bioremediation

Oil spills in refinery, shore sand and processing factories could be handled by the

use of lipases from different origins (Demarche et al., 2011). It has been also used for the

degradation of wastewater contaminants such as olive oil from oil mills. Another important

application has been reported for the degradation of polyester waste, removal of biofilm

deposits from cooling water systems and also to purify the waste gases from factories

(Anonymous, 1995).

2.7.6 Detergency and cleaning

An important application of lipases resistant to high temperatures, proteolysis, and

denaturation by surfactants, is their use in laundry detergents along with proteases to

improve the removal of lipid stains. They are also used in the synthesis of surfactants for

soaps, shampoos and dairy products (Hasan et al., 2006; Horchani et al., 2009; Pandey et

al., 1999; Schmid and Verger, 1998).

2.7.7 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 (Dubé et al., 2008;

Hasan et al., 2006).

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2.7.8 Other applications

Lipases are also used in biodiesel production, leather processing, hard-surface

cleaning, single-cell protein production and so on (Hasan et al., 2006; Horchani et al., 2010;

Kademi et al., 2006; Tan et al., 2010).