PRODUCTION AND CHARACTERISATION OF LIPASE FROM ASPERGILLUS NIGER IN A SUBMERGED FERMENTATION SYSTEM

ABSTRACT

Three fungal isolates (FSP1, FSP2 and FSP3) were isolated based on morphological characteristics from laundry waste water disposal site. A 10 day pilot study on the three isolates in submerged fermentation system using olive oil as sole carbon source revealed that FSP1 had the highest lipase production on day 9 and pH 4.0 was the optimum pH for the enzyme production. Molecular studies on the ribosomal DNA of FSP1 showed an internally transcribed spacer (ITS) region of 550-600 base pairs. Phylogenetic analysis of the ITS sequence identified the organism as Aspergillus niger and the organism was used for enzyme production. Ammonium sulphate saturation (70%) precipitated protein with the highest lipase activity. After dialysis and gel filtration (Sephadex G-

100), the active fractions were pooled together and characterised. The protein concentration and the specific activity of the crude enzyme were 1.685 mg/ml and 121.721 U/mg, respectively. After the partial purification process, a 10-fold purification was obtained with specific activity of

1185.05 U/mg. The pH and temperature optima for the enzyme activity were 5.5 and 50 C, respectively, using p-nitrophenyl palmitate (pNPP) as substrate. The Michealis-Menten constant, Km and maximum velocity, Vmax were 70.79 mg/ml and 227.27 µmol/min, respectively. Calcium ion (Ca2+) and cobalt ion (Co2+) enhanced lipase activity in a concentration dependent manner, whereas Mn2+ and Fe2+ inhibited lipase activity. The stability studies revealed that at pH 6.0, the enzyme retained more than 60 % of its activity for two hours. Also, at temperature range of 40–50 °C, the enzyme maintained more than 60% of its activity for two hours. The least thermal denaturation constant, kd obtained at 40 C was 0.0255 min−1. The enzyme also have maximum D– value (99.29 min) at 40 C. The half-life of inactivation (t1/2) decreased with successive temperature increase with 80 C having the least half-life of 1.8 min. The Z-value which represents the enzyme’s sensitivity to heat treatment was 37 ºC. The activation energy of inactivation (Ea(inact.)) was 58.097 KJmol–1K–1. The thermodynamic parameters for inactivation appeared to be temperature dependent. The change in enthalpy of inactivation (ΔH) decreased with increasing incubation temperature and had its least value of 55.163 KJ/mol at 80 C and maximum value of 55.495 KJ/mol at 40 C. Similarly, the maximum free energy change (ΔG) of inactivation (52.92 KJ/mol) was obtained at 40 C. The mean entropy change (ΔS) of inactivation was 0.009 KJmol– 1K–1. These results revealed that the lipase  was sensitive to heat treatment and its thermal inactivation was

CHAPTER ONE

INTRODUCTION

The exploitation of enzymes by man have transcended ages. Enzymes have been used in various forms and medium especially as extracts obtained from microorganisms, plants or animal organs. The potentials derivable from the use of enzyme technology led to extensive research on enzyme sources and their applications. Successful commercial exploitation of enzymes firmly established industrial biotechnology sector (Raveendran et al., 2018). The blooming industrial enzyme market became a major revenue earner in industrial science sector as large number of enzymes are produced, sold and put into numerous uses (Li et al., 2012). The major industrial enzymes include carbohydrases, proteases and lipases which act on carbohydrates, proteins and lipids respectively (Sangeetha et al., 2011).

Lipids are one of the most abundant macromolecules that play crucial physiological roles and lipolytic enzymes are required for their metabolism (Barros et al., 2010). Lipolytic enzymes include esterases commonly called carboxyesterases (EC. 3.1.1.1) and lipases (triacylglycerol acylhydrolase, EC. 3.1.1.3). The basic difference between esterases and lipases is that the former acts on short chain triglycerides which are soluble in water. Lipases, however, act on lipids which form aggregates in water and require a lipid-water interface for its catalysis (Treichel et al., 2010). They hydrolyze long chain water-insoluble triglycerides into their component diglycerides, monoglycerides,  glycerol  and fatty  acids.  However in  micro-aqueous  environment,  they can catalyse fatty acid esterification, trans-esterification, aminolysis, and acidolysis (Hassan et al.,

2013). Many industrial applications of lipase focus on its regiospecificity and enantiospecificity

(Brahmachari, 2017).

Lipases have been isolated from a large number of plant, animal and microbial sources (Ramnath et al., 2017b). The ease, with which enzymes could be isolated from microbes, has made both bacteria and fungi as predominant sources of lipase. In addition, they can be produced through solid state or submerged fermentation techniques with less space and time requirement, as well as easy opportunity for process modification and optimization (Ranveendran et al., 2018).

Generally, lipases do not require cofactors to hydrolyze there substrates and remain active in the presence of organic solvents. These properties open numerous vistas for industrial application of lipase and put the enzyme on the front burner of research (Barros et al., 2010). However, the outstanding limitation in application of lipases is the unavailability of lipases possessing all the specific required industrial characteristics. Moreover, the growing demand for lipases create the need to source for microorganisms capable of producing efficient and cost-effective lipases using cheap carbon and nitrogen sources (Lubertozzi and Keasling, 2009). The lipases so produced are further characterized for pH, temperature, substrate specific, stability and bioenergetics. This will enable the prediction of suitability of the enzyme for industrial application (Ramnath et al., 2017a).

Aspergillus species are among the organism of great significance in microbial enzyme production and enzyme technology. The organism is versatile and flexible in its carbon and nitrogen source requirement. Moreover, most of the species are considered safe and the enzymes they produced are inducible and extracellular, thus making the organism an easy, efficient and cost-effective means for industrial enzyme production (Schuster et al., 2002; Lubertozzi and Keasling, 2009).

1.1       Enzymes

Enzymes have become an essential constituent of several industrial processes. They have found application in food, beverage, textile, detergent, pulp and paper, medical and pharmaceutical industries (Ramnath et al., 2017a). In all these industry, enzymes act as biocatalysts where they catalyze the conversion of substrates to the desired products. The enzyme revolution in industrial process have basis in their unique ability to lower the energy barrier of chemical reactions, producing outstanding increase in reaction rates up to over a billion fold (Gurung et al., 2013). Sangeetha et al. (2011) identified carbohydrases, proteases and lipases which act on carbohydrates, proteins and lipids respectively as major enzymes of industrial importance. These three groups of enzymes contribute an estimated 70% of the global enzyme sales (Li et al., 2012). Table 1.1 shows various industries, the enzymes used and their application in the in such industries.

Table 1.1: Enzymes used in various industrial processes and their application

Source: Ramnath et al., (2017a)

Application

	Amylase Lipase	Starch stain removal Lipid stain removal
	Cellulase	Cleaning, colour clarification
Fuel	Xylanase Lipase	Viscosity reduction Synthesis of lipase-catalyzed biodiesel
Food	Protease Lipase Lactase	Milk clotting, flavour Improvement of food texture Lactose removal (milk)
	Pectin Methyl esterase Pectinase Transglutaminase	Firming fruit based products Fruit-based products Modify visco-elastic properties
Baking	Amylase Xylanase	Bread softness and volume Dough conditioning
	Lipase Phospholipase Glucose oxidase Lipoxygenase Protease Transglutaminase	Dough stability and conditioning Dough stability and conditioning Dough strengthening Dough strengthening, bread whitening Biscuits, cookies Laminated dough strengths
Pulp and paper	Lipase Protease Amylase Xylanase Cellulase	Pitch control, contaminant control Biofilm removal Starch coating, deinking, drainage improvement Bleach boasting Deinking, drainage improvement, fiber
		modification
Fats and oils	Lipase Phospholipase	Transesterification Degumming, lysolecithin production
Organic synthesis	Lipase Acylase Nitrilase	Resolution of chiral alcohols and amides Synthesis of semisynthetic penicillin Synthesis of enatiopure carboxylic acid
Leather	Protease Lipase	Unhearing, bating Depickling
Environmental application	Lipase	Removal of solid and water pollution by hydrocarbons, oils and lipids

Protein stain removal

Apart from the unique high reaction rates offered by enzymes, they also offer other important advantages. There are characteristically resilient enzymes that can withstand harsh industrial processes such as extreme pH ranges, high temperatures, and the presence of surfactants and organic solvents. They also display distinctive chemospecificity, stereospecificity, and regioselectivity that are of particular interest in applications, such as the synthesis of optically pure compounds (Haki and Rakshit, 2003). Enzymes are sourced from animal, plants and microbial

origin. However, microbial sources are most preferred due to their high yield, easy, consistent and cost-effective production. They are also found to possess better stability, selectivity, and broad substrate specificity. In addition, they can be produced through fermentation techniques with less space and time requirement, with easy opportunity for process modification and optimization (Raveendran et al., 2018).

1.2       Lipases

Lipases (triacylglycerol acylhydrolases, EC 3.1.1.3) are enzymes which catalyze the hydrolysis of long chain triglycerides. They are naturally present in the stomach and pancreas of humans and other animal species in order to digest fats and lipids. They are also resident in blood, intestinal juices, and  adipose tissues.  Lipases  catalyse the hydrolysis  of triglycerides  (fats)  into  their component fatty acid and glycerol molecules. Under suitable conditions, lipases can catalyse fatty acid esterification, trans-esterification, aminolysis, and acidolysis reactions (Raveendran et al.,

2018).

1.2.1    History of Lipase

Lipase was  reported  to be first discovered in  1825 by Claude Bernard in pancreatic juice. Subsequently, extracts of animal pancreas became conventionally used as the source of lipase for commercial applications (Sangeetha et al., 2011). However, with the establishment of numerous industrial potentials of lipase, occasioned by increased use of organic esters in biotechnology and the chemical industry, the demand for lipase could not be met from animal sources (Stergiou et al., 2013). Microbial sources of lipase became the major focus and it proved to be cost effective, reliable, and have robust applications (Raveendran et al., 2018)

1.2.2    Reactions Catalyzed by Lipases

Lipases catalyse the hydrolysis of long chain triacylglycerol substrates. The acyl group of the triglyceride are made of carbon chain greater than eight (> C8). This is in contrast to esterases that catalyse water soluble triglyceride having acyl carbon chain less than or equal to eight (Bornscheuer, 2002).

Figure 1.1: Hydrolysis and synthesis of triacylglycerol by lipase (Messaoudi et al., 2010).

The reactions catalysed by lipase could broadly divided into three: hydrolysis, organic synthesis, and aminolysis. Lipases, as hydrolyzing agents are active in environments, which contain a minimum of two distinct phases, where all reactants are partitioned between these phases (Stergiou et al., 2013).

Figure 1.2: Hydrolysis reaction of lipase (Barros et al., 2010)

Organic synthesis reactions catalysed by lipase such esterification, transesterification, and regioselective acylation require restricted water environment in order to occur (Reis et al., 2009). Esterification reactions of lipase stands out as the most significant chemical and biochemical processes of industrial relevance due to an increased use of organic esters in biotechnology and the chemical industry.

Figure 1.3: Esterification reaction of lipase (Barros et al., 2010)

Transesterification could be acidolysis (exchange of groups between and ester and an acid), acoholysis (exchange of groups between an ester and an alcohol), or interesterification (exchange between two esters) (Barros et al., 2010).

Figure 1.4: Transesterification reactions of lipase

In aminolysis, water, the natural nucleophile is replaced by an alcohol, hydrogen peroxide or amine

(Fernandes et al., 2004).

Figure 1.5: Aminolysis reaction of lipase (Barros et al., 2010)

1.2.3    Classification of Lipases

Lipases have been classified based on a number of criteria. Based on their sources, they have been classified as animal, plant, microbial and recombinant lipases (Sangeetha et al., 2011). They could be further classified as acidic, neutral or alkaline lipases based on their pH optima and pH stability (Sharma et al., 2016).   Based on their temperature optima and thermal stability, lipases are classified as psychrophilic, mesophilic and thermophilic lipases (Sharma et al., 2016). Bacteria lipase which remains the most studied of microbial lipases are classified into eight families (families I – VIII) based on differences in their amino-acid sequences, the conserved motifs and

biological properties (Arpigny and Jaeger, 1999). The classification enables the prediction of: important structural features such as residues forming the catalytic site or the presence of disulphide bonds; types of secretion mechanism and requirement for lipase-specific foldases; and the potential relationship to other enzyme families (Arpigny and Jaeger, 1999, Ramnath et al.,

2017a).

1.2.4    Protein Chemistry of Lipase

A diverse array of genetically distinct lipases are found in nature and as such are always found to possess varying length of amino acid residues. The range of molecular weight of most microbial lipases are between 20kDa and 75kDa (Messaoudi et al., 2010). The secondary structure consisting alpha helix and beta sheets are cast into several types of protein folds (Natalello et al., 2005). However, the catalytic mechanisms are mostly built on an alpha/beta hydrolase fold which employ a chymotrypsin-like hydrolysis mechanism (Ramnath et al., 2017a).

1.2.4.1 Primary Structure

The molecular weight of most microbial lipases vary between 20kDa and 60 kDa due to the differences in length of amino acid residues of lipase from different sources (Mala and Takeuchi,

2008). Though, microbial lipases of larger molecular weight range of 65-75 kDa has been reported (Arpigny and Jaeger, 1999). Lipases require no cofactor and as such are non metal containing enzymes. However, some lipases have been reported to possess two aspartate residues involved in Ca2+-binding. The localization of these residues within the vicinity of the catalytic histidine and aspartate residues suggests that they are important in the stabilization of the active site of these enzymes (Sangeetha et al., 2011). Lipase has a discrete distribution of hydrophobic and hydrophilic residues. Hydrophobic collapse contribute to much of the secondary and tertiary structures, as the hydrophobic core residues make up the interior of the protein, while polar residues are on the surface (Khan  et al., 2017).

Most lipase contain a conserved pentapeptide region which consist of Gly-Xaa-Ser-Xaa-Gly. The residue Xaa could be any amino acid. The glycine residues are sometimes substituted by small amino acid residues such as alanine. The serine residue located in this conserved region acts as the nucleophile in the catalytic triad of Ser, His, and Asp (Glu) (Ramnath et al., 2017a).

1.2.4.2 Secondary Structure

Lipases are built on α-helixes stabilized by β-sheets forming α/β hydrolase structure. The active site which contains the catalytic triad is covered by a lid. The lid is made of α-helical structure (Mala and Takeuchi, 2008). The rotation of the lid is crucial to interfacial activation of lipases. Also, the number of helices forming the lid domain as well as the position of the lid has a direct correlation to the optimum reaction temperature and thermal stability of lipases (Khan et al., 2017). In Candida rugosa the α-helix lid covering the active site is formed by residues 65-95. The lid structure is stabilized by disulphide bond between Cys60-Cys97 (Akoh et al., 2004; Colton et al.,

2011). In addition Fourier transform infrared spectroscopy study suggests the existence 40% α- helices, 23% β-sheets, while the turns constitute 28%. The α-helices exist in two forms with peaks at 1657.9cm−1 and 1649.4cm−1, while the β-sheets also exist in two forms with peaks at 1630.0cm−1 and 1637.2cm−1 (Natalello et al., 2005).

1.2.4.3 Three Dimensional Structure of Lipase

The three dimensional (3D) structures of lipase determine their regiospecificity and enantiospecificity essential for suitability of lipases for specific applications. The 3D structure of microbial lipases have been the most studied (Mala and Takeuchi, 2008).

Figure 1.6: Crystal structure of Geobacillus zalihae lipase with two helices in its lid domain. Catalytic triad residues were highlighted by yellow sticks and lid domains were highlighted by blue colour (Matsumura et al., 2008)

In spite of amino acid dissimilarity among lipases from difference sources, they have very similar folds. Hence, lipases have comparable three-dimensional structure characterized by α/β-hydrolase folding which is a specific sequence of β-strands connected by α-helices (Khan et al., 2017).

The canonical α/β-hydrolase fold consists of a central parallel β sheet of eight strands with the second strand anti-parallel. The parallel strands range are connected by α helices which pack on either side of the central β sheet. The β sheet has a left-handed super-helical twist such that the surface of the sheet covers about half a cylinder and the first and last strands cross each other at an angle of 90. The curvature of the β sheet may differ significantly among the various enzymes and also, the spatial positions of topologically equivalent α helices may vary considerably. They differ substantially in length and architecture, in agreement with the large substrate diversity of these enzymes (Jaeger et al, 1999; Andualema and Gessesse, 2012).

1.2.4.4 Quaternary Structure of Lipase

The multimeric form of the enzyme has its monomers held together by disulphide bonds (between cysteine residues), hydrogen bonds (between main chain and side chain atoms), and electrostatic bonds (between positively charged nitrogens of Arginine and Lysine, with the negative charge of oxygen atoms of Aspartate and Glutamate. Pancreatic lipase has twelve disulphide bonds (Protopedia, 2018).

1.2.4.5 Active site of Lipase

At the core of the lipase is a helical segment called the lid that covers the active site when the enzyme is in closed conformation. The lid blocks solvent from accessing the active site thereby protecting the oxyanion hole. The opening of the lid arises from alteration of the secondary structure of lipase binding site from a closed to an open ring structure. This happens when lipid aggregates  are  available,  the  lid  opens  immediately  and  the  enzyme  assumes  the  open

conformation thereby increasing its activity. This process is known as interfacial activation

(Thomas et al., 2005).

The active site of lipases normally contain the catalytic triad typical of all α/β-hydrolase fold enzymes. The catalytic triad is comprised of Ser-Asp-His (Glu instead of Asp for some lipases). There is also a pentapeptide consensus sequence (Gly-x-Ser-x-Gly) found around the serine of the active site (Ramnath et al., 2017b). The serine residue act as the nucleophile, while the Asp (Glu) residue is the catalytic residue. The nucleophilic Ser residue is located at the C-terminal end of β strand in the highly conserved pentapeptide, forming a characteristic β-turn-α motif which is known as nucleophilic elbow (Andualema and Gessesse, 2012).

With the exception of catalytic triad, most residues in the active site are hydrophobic, and their main chain atoms are not accessible for hydrogen bonding. The hydrophobicity is necessary for hydrophobic interaction with the substrate – the alkyl chain of a triglyceride (Khan et al., 2017).

The hydrolysis of the substrate commence with a nucleophilic attack by the oxygen atom of the active-site serine on the carbonyl carbon atom of the ester bond, leading to the formation of a tetrahedral intermediate stabilized by hydrogen bonding to nitrogen atoms of main chain residues that belong to the oxyanion hole (Khan et al., 2017). An alcohol is produced and released from an acyl-lipase complex which is finally hydrolyzed with the production of the fatty acid and regeneration of the enzyme (Andualema and Gessesse, 2012).

Figure 1.7: Hydrolysis of substrate at the active site of lipase (Protopedia, 2018). (a) acylation of the enzyme, (b) deacylation of the enzyme-acyl complex.

1.2.5    Lipase Screening and Production Media

1.2.5.1 Screening and Assay

Several methods have been proposed for screening of lipase production. These methods either directly use the microorganism under study (Gopinath et al., 2013) or measure lipolytic activity in the crude or purified culture preparations (Singh et al., 2010). The plate detection methods use agar plates containing lipid substrate such as Tween 20 and tributyrin, and lipolysis was observed as clear halos or opaque zones around the well containing culture or enzyme preparation. However, the observed zones of clearance could be as a result of actions of esterases when tributyrin is used (Gopinath et al., 2013). Plates containing chromogenic substrates with pH indicators like phenol red or Victoria blue are also used. A decrease in pH due to release of fatty acids due to lipolysis causes change in the colour of the indicators is measured. Agar plates supplemented with Rhodamine B which indicate lipolysis by the formation of fluorescent orange halos are widely used due to their sensitivity and reliability. The colorimetric methods are based on measuring the complexes formed between the released free fatty acids and a divalent metal ion, usually copper (Gopinath et al., 2013). The widely adopted spectrophotometric methods use designed substrates usually p-nitrophenyl esters of fatty acids and the lipase activity is assayed by measuring the amount of p-nitrophenol released after enzymatic hydrolysis of the ester (Wang et al., 2009). The esters used range from the two carbon esters p-nitrophenyl acetate to eighteen carbon p-nitrophenyl stearate (Ramnath et al., 2017b). The p-nitrophenyl esters of longer fatty acid length are more specific for lipase assay. This is because esterases can also hydrolyse short carbon chain fatty acid esters (Sangeetha et al., 2011). The titrimetric method employs the neutralisation of the free fatty acids that are released after lipolysis and the volume of the base consumed indicates the extent of lipolysis. The other least commonly used methods like chromatographic, turbidimetric, fluorimetric, immunological and radioactive assays (Hasan et al., 2009).

1.2.5.2 Fermentation Systems for Lipase Production

Production of microbial lipases have been reported to be done by both submerged and solid state fermentation methods (Sarkar and Laha, 2013). Fermentation is the procedure of biological conversion of complex substrates into simple compounds by various microorganisms. It has been extensively applied for the production of many microbial enzymes (Liu and Kokare, 2017).  Each of these fermentation methods seems to present its own unique advantages and disadvantages. The solid state fermentation is an attractive option for microbial enzyme production due to the prospect of using waste or by-products of agricultural origin as both support for microbial development and as its nutrient source. This approach in lipase production offers the opportunity to create high value from low cost substrate resulting in reduction of the cost of enzyme produced and recycling of agro waste (Liu and Kokare, 2017). On the down side, the scale up production of lipase through solid state fermentation project difficulties due to uneven distribution growth conditions across the solid substrates. While the large scale submerged fermentation require higher capital for equipment and control systems, and recurring expenditure for its complex media. Despite the vistas of opportunities in solid state fermentation process, most microbial lipases are produced mostly by submerged culture. This is because submerged fermentation offer greater control of growth conditions than solid state fermentation (Sangeetha et al., 2011).

Another method for lipase production is the use of immobilized cell cultures. This method offers the dual advantage of reuse of immobilized microbial cells while at the same time smoothening the downstream processing. The major drawback is the cost and complexity of immobilization process (Gunasekaran and Das, 2005).

1.2.5.3 Media development

The aim of microbial production of lipase is to obtain a high extracellular yield of the enzyme. Lipase production is subject to the type and concentration of nitrogen and carbon source. It is also influenced by the production conditions such as the temperature, pH and oxygen concentration

(Ertugrul et al., 2007). Studies on optimal culture and nutritional requirement for lipase production shows that carbon sources with high lipid content are essential to high yield of lipase. Thus, a lot of studies on lipase production use lipid substrates as sole carbon source as against the use of simple sugars (Zhang et al., 2009). Though, good lipase yield have also been obtained in absence of fats and oil. In some other cases, simple sugar is used as carbon sources and lipid substrates are added as inducers (Hun et al., 2003).

Extracellular production of lipases appears to be non-constitutive. Hence the use of inducers such as Tween 20 and Tween 80, vegetable oils, synthetic triglycerides like tributyrin and tripalmitin, and hexadecane (Sangeetha et al., 2011).

Surfactants are another essential media component required for optimum lipase production. The presence of surfactant is an important pre-requisite for maximum lipase production. Surfactants reduce the surface tension between aqueous and organic phase present in lipase production media. This formation of emulsion known as interfacial activation presents an interface for enzyme activity. Frequently used surfactants include Triton-X 100 and Tween (Pogaku et al., 2010).

Generally, the presence of surfactants boosts the lipid-water interface thereby enhancing the rate of lipolysis. However, this assertion is not always the case for all surfactants. While Tween-80 and Triton X-100 have been reported to have stimulatory effect on lipase production and activity, SDS produces the opposite effect. In addition, the effect of surfactants is dependent on the concentration used. For instance, using Baccillus pumilus to produce lipase at varying concentration of Tween-

80, the optimum enzyme production was found to be at 0.5% concentration of Tween-80 while concentration of the surfactant up to 1% inhibited lipase production (Zhang et al., 2009).

Nitrogen source is another important nutritional requirement for microbial lipase production. Peptone is among the nitrogen sources reported to augment lipase production. In contrast to carbon sources, nitrogen source have not been reported to suppress lipase synthesis (Gunasekaran et al., 2006).

1.2.6    Properties of Lipase

The industrial applications of lipases is a function of its unique characteristics. Some properties of lipase vary from one source to the other. Such factors are pH and temperature optima, tolerance of

emulsification and surfactants, temperature and pH stability, substrate specificity, and storage capability are important considerations in the selection and development of a commercially useful product (Liu and Kokare, 2017). Other properties such as the ability to utilize comparatively wide spectrum of substrates, none requirement of cofactors and reaction specificity (enantioselectivity and regioselectivity) are of remarkable interest to industrialists and researchers (Saxena et al.,

2003).

1.2.6.1 Determination of Lipase Activity

Determination of most of these characteristics of lipase is achieved by subjecting the enzyme to the condition under study, after which the activity of the enzyme is determined based on the hydrolysis of p-nitrophenylesters of fatty acids with various chain lengths. This is determined by spectrophotometric detection of p-nitrophenol at 410 nm. For synthetic reaction, lipase activity is determined by the speed of ester formation using gas chromatography (Fernandes et al., 2004). Enantioselectivity is determined using gas chromatography or High Performance Liquid Chromatography (HPLC), with chirally modified columns (Andualema and Gessesse, 2012).

1.2.6.2 pH Optima

Bacterial lipases mostly have their pH optima in the alkaline region, making the valuable catalysts for numerous industrial processes (Ahmed et al., 2009). The fewer acidic lipases are of fungal origin. Though the bacteria Pseudomonas gessardi was reported to have pH optima in the acidic region (Ramani et al., 2010). Studies on lipases from Candida cylindracea, Humicola lanuginosa and Thermomyces lanuginosa suggests that the type of emulsion system as well as the substrate been hydrolyzed appears to have an effect on the pH optima of lipases (Fernandes et al., 2004).

1.2.6.2 Temperature Optima

Conventional microbial lipases have optimum activity at the temperature ranges between 30C and 60C. This is due to high entropy and increased fluidity of interface of the two phases (the aqueous and organic phases) required for lipase activity, leading to faster inactivation and low ability to detect activity after microencapsulation (Fernandes et al., 2004). The exceptions to this

temperature range are lipases obtained from extremophiles. Psychrophilic lipases from Antarctic Pseudomonas and Moraxella sp have peak activity at low temperature while the activity of lipase from the thermophile Bacillus thermocatenulatus peaks 70°C (Fernandes et al., 2004). These extreme and remarkable features offer the prospect of using the enzyme in reactions carried out at low or high temperature (Cai et al., 2009).

1.2.6.3 Effects of Substrate Concentration

The effects of substrate concentration on enzymes are studied using parameters such as the Km and Vmax of the enzyme. These parameters are usually determined for enzymes that follow Michaelis- Menten curve. The Km is a measure of affinity of the enzyme for the substrate and the relationship of Km to affinity is inverse. For lipase from Thermomyces lanuginosa, the Km value of the enzyme using the substrates triolein and tributyrin was found to be 150mM and 120mM respectively, while their respective Vmax is as 1935Umg−1 and 485Umg−1. In addition, the effect of substrate concentration  follows  the Michaelis  curve except  in  the case of triolein  where the activity decreased as substrate concentration beyond 200mM. This could be caused by the direct influence of the high concentration fatty acid chains of the substrates on the lipase which alters the micellar structure (Fernandes et al., 2004).

1.2.6.4 pH Stability

Lipase from Thermomyces lanuginosa proves to be very stable as it remains active over a pH range of 7.0 to 11.0 for one hour (Fernandes et al., 2004). Lipase isolated from from Bacillus stearothermophilus remained stable at a pH range of 7–9 after incubation for one hour at 30°C, with a residual activity remaining above 50% (Massadeh and Sabra, 2011). The extremophile Geobacillus thermodenitrificans produce lipase found to be most stable at pH 7.0 retaining its initial activity for a three hour duration at its optimal temperature of 65°C (Balan et al., 2012). For Pseudomonas aeruginosa, the enzyme was found to be stable for two hours at pH 6.9 (Borkar et al., 2009). In the case of lipase from Pseudomonas fluorescens the enzyme was found to retain 100% of its activity for a period of 45mins and reduced to 70% after 100mins at pH 8.0 (Gökbulut

and Arslanoğlu, 2013).

1.2.6.5 Thermostability

Thermostability is the product of the protein’s amino acid sequence, which provides conformational rigidity to the enzyme through intramolecular interactions, with the internalization of hydrophobic residues and superficial exposure of hydrophilic residues (Colla et al., 2015). Thermostability is usually expressed in terms kinetic parameters: denaturation rate constant, half- life and thermal death concepts (D-value and Z-value); while thermodynamic parameters is used to express the of structure-stability relationship (Borba et al., 2018).

Based on thermal stability, lipases could be classified as psychrophilic, mesophilic or thermophilic. Thermostable lipases from many Pseudomonas and Bacillus sp. have been isolated and studied. Mesophilic and thermophilic organisms produce more stable lipases than psychrophiles (Ahmed et al., 2009). Lipases from extremophiles (psychrophiles/cryophiles and thermophiles) prove to be more valuable for biotechnological applications due to their stability at extremes of temperature (Haki and Rakshit, 2003). High temperature stable lipases have been isolated from Bacillus species (Timucin and Sezerman, 2013) and Pseudomonas sp (Yang et al., 2015).

Mala and Takeuchi (2008) showed a clear relationship between lipase thermal stability and the enzyme structure. Other factors such as metal ions, the number of hydrogen bonds, salt bridges, stabilization of secondary structures, occurrence of disulfide bonds, higher number of proline residues, higher polar surface area, shortening of loops, and stabilization of the lid domain (Khan et al., 2017).

Psychrophilic lipases are commonly found among cold adapted microorganisms. Some organisms that produce cold active lipases include Aeromonas sp., Rhizopus sp. and Mucor sp, Pseudoalteromonas sp. and Psychrobacter sp., Photobacterium lipolyticum and Geotrichum sp. The interest in lipase having high activity at low temperature is due to several industrial applications to which it could be put to. They are used in synthesis of delicate chiral organic chemicals or pharmacological importance; as food additives for fermentation, cheese manufacture, and meat tenderizing; and as additives in detergent for cold washing (Cai et al., 2009; Andualema and Gessesse, 2012).

1.2.6.6 Effect of Metal ions on Lipase

Metal ions augment the catalytic activity of enzymes and also bestow thermostability to them. The presence of metal ions is crucial for the maintenance of active sites of numerous enzymes.

Moreover, some enzyme uses these metal ions as cofactors in their catalytic activity (Sauza et al.,

2014). Interestingly, lipases from diverse sources respond differently to different metal ions. The commonly studied metal ions are Ca2+, Zn2+, Mg2+, Mn2+, Co2+, Hg2+, Cu2+, and Fe2+. While some metal ions have stimulatory effect on some lipases, the same metal ions also inhibit other lipases (Ahmed et al., 2009). Despite the conflicting reports on effects of metal ions on lipases activity, Ca2+ has been found to be demonstrate a stimulatory effect on all lipases that are found to be sensitive to metal ions (Ahmed et al., 2009; Zhang et al., 2009). In these lipases, the folded enzyme contains a region rich in negatively charged amino acids. Electrostatic repulsion around this region is detrimental to enzyme stability. The binding of Ca2+ at this negatively charged region imposes structural alterations. The Ca2+ ion forms a bridge analogous to disulphide bond which crosslink the polypeptide chain. This offers stability and rigidity to the enzyme (Sangeetha et al., 2011. The presence of Ca2+ binding site can be inferred from inactivation of lipase by EDTA. This inactivation is relieved by addition of CaCl2. Importantly, lipases which exhibit Ca2+ independent thermostability and activity have been reported by Kim et al. (2002). These lipases function effectively in the presence of EDTA usually found in laundry detergents, thereby making the enzyme priceless in laundry detergents (Kim et al., 2002).

1.2.6.7 Tolerance to Organic Solvents

Organic solvents are preferred to their inorganic counterparts in bio-reaction systems. This is because organic solvents offer increased substrate solubility, easy enzyme and product recovery and helps in driving the equilibrium in the forward direction for synthetic reactions (Zhang et al.,

2009). Lipases that are tolerant to organic solvent find applications in trans-esterification reactions, synthesis of biopolymers and production of biodiesel (Singh et al., 2010). The stability and activity of lipase is usually assessed in the presence of organic solvents such as isopropanol, n-hexane, n- heptane, n-octane, n-decane, benzene, methanol, ethanol, chloroform, acetic acid, acetone, glycerol, toluene, xylene, styrene, benzene, ethylbenzene, cyclohexane, dimethylsulfoxide and tetrahydrofuran (Zhang et al., 2009). Lipases from different sources display different degrees of tolerance to various organic solvents and thus commercial utilization of lipase requires an in-depth analysis of its tolerance to these solvents. The sensitivity of lipases to solvents varies in accordance with the extent of the solvent’s polarity. Hence, polar solvents have more destabilizing effect than non-polar solvents (Ahmed et al., 2010).

1.2.6.8 Substrate Specificity

Specificity of lipases is influenced by an array of properties such as substrate type, substrate stereochemistry, positions of the fatty acid ester and molecular properties of the enzyme. These factors alter the binding and substrate preferences of the enzyme (Ramnath et al., 2017b).

By basic definition, lipases in liquid medium, catalyze the hydrolysis of ester bonds present in acylglycerols releasing free fatty acids and glycerol. Lipases take effect on ester bonds present in acylglycerols to release free fatty acids and glycerol in a liquid medium. In liquid controlled environments, these enzymes have the capacity to carry out the reverse synthetic reaction (esterification) through acidolysis, interesterification, and alcoholysis (Hassan et al., 2013).

Based on specificity and reaction catalysed, lipases according to Barros et al. (2010) could be grouped as:

1.         Non substrate selective lipases: These have glycerol esters as their natural substrates and hydrolyze triacylglycerols, diacylglycerols, monoacylglycerols and phospholipids.

2.         Regioselective lipases: These are subdivided into: (i) Non-regiospecific lipase that catalyze complete hydrolysis of triacylglycerols through mono and diacylglycerol intermediates into glycerol and fatty acids in a random manner; and (ii)  Specific 1, 3 lipases: these only hydrolyse triacylglycerols at the C-1 and C-3 glycerol bonds, producing fatty acids, 2-monoacylglycerols and 1,2 or 2,3-diacylglycerols, the latter two being chemically unstable, promoting migration of the acyl group producing 1.3-diacylglycerol and 1 or 3-monoacylglycerols.

3.         Fatty acid selective lipases: these can be specific for a specific type of fatty acid or a particular group of fatty acids. They hydrolyze the fatty acid esters independent of their position on the glycerol backbone. An example is Canola seed lipase which discriminates fatty acids with cis-4 or cis-6 double-bonds.

4.         Enantioselective lipases: These have the ability to discriminate enantiomers in a racemic mixture. An example is the R-isomer of sparteine 9, which tastes sweet, whereas the S-isomer tastes bitter. The enantiospecificities of lipases can vary according to the substrate and this variation can be connected to the chemical nature of the ester.

Figure 1.8: Reactions catalyzed by non-specific and 1,3 specific lipases (Barros et al., 2010)

1.2.7    Sources of Lipase

Lipases are ubiquitous in nature. They could be sourced from animal, plant or microbial origin. Lipases of microbial origin are the most widely used source of the enzyme for biotechnological applications and organic chemistry (Ranveendran et al., 2018).

1.2.7.1 Animal lipases

These are the first source of lipase to be exploited. They have been isolated from insects, fishes, mammals. They are found localized in their secretory organs or at the organs where they perform their lipolytic functions. These lipases play a vital role in digestion, storage and mobilization of lipids in biological systems. The availability of other lipase sources, especially microbial lipases makes animal lipases the least studied. The major animal lipases are pancreatic lipase, gastric lipase, intestinal lipase and lipases of the adipose tissues (Ranveendran et al., 2018).

1.2.7.2 Plant lipases

In plants, lipases are present in the food reserve tissues of growing seedlings, especially those which possess a large amount of triacylglycerols. Lipase activity in plant seeds peaks during germination because the triacylglycerols are converted to soluble sugars by the action of lipase which is then transported to the growing tissues to supply structural carbon and energy, required

to provide support for the growth of young plants. The optimum pH range of most plant lipases is between 4.0-8.0, while their temperature optima varies from 25-60C (Barros et al., 2010).

1.2.7.3 Microbial Lipases

Lipases of microbial origin represent the most studied, characterized and applied class of the enzyme. They find application in organic chemistry, food and industrial biotechnology. Microbial lipase production varies according to the microorganism, the composition of the growth medium, cultivation conditions, pH, temperature, and the type of carbon and nitrogen sources (Sangeetha et al., 2011; Brahmachari, 2017).

1.2.7.3.1 Bacteria lipase

Compared to other sources of lipase, bacteria lipases are most studied. Bacterial lipases are mostly glycoprotein, though some extracellular bacterial lipases are lipoprotein and are the commercially important lipases due to their yield and ease of production. Lipases from Pseudomonas and Bacillus were probably the first studied and have preponderant role in industries. Other bacteria genera exploited for lipase production are Achromobacter, Alcaligenes, Staphylococcus, Burkholderia, Chromobacterium, Enterococcus, and Corynebacterium (Thakur, 2012). Most bacteria lipases that has been characterized for pH, temperature appears to possess optimum activity at pH ranges from 4.0-10.0 and temperature ranges from 27-80C (Khan et al., 2017).

1.2.7.3.2 Fungal lipase

Most commercially important lipase-producing fungi are recognized as belonging to the genera Rhizopus, Candida, Aspergillus, Penicillium, Geotrichum, Humicola, Mucor, and Rhizomucor. Lipase production by fungi varies according to the strain, the composition of the growth medium, cultivation conditions, pH, temperature, and the type of carbon and nitrogen sources. The industrial demand for new sources of lipases with different catalytic characteristics stimulates the isolation and selection of new strains. Lipase-producing fungi have been found in different habitats such as industrial wastes, vegetable oil processing factories, dairy plants, and deteriorated food (Thakur,

2012).

1.2.8    Application of Lipases

Lipases have found application in numerous industry using biocatalysts in their production. They are extensively applied in food industry, medical and pharmaceutical industries, textile industry, biofuel, detergents, animal feed, leather, and paper processing (Gerits et al., 2014; Raveendran et al., 2018)

1.2.8.1 Lipases in Textile Industry

In the textile industry lipases are used for desizing of fabrics. Desizing is the process of removing size materials infused into fabric during weaving. Cellulases, amylases, proteases and lipases are the enzymes used depending on the sizing agent. They replace previously used acids and oxidizing agents which damage cellulose materials of fabric. Also the enzymes improve fabric’s absorbance thereby enabling smoothness and consistency dyeing. This is very important in polyester materials known for their strength and wrinkle resistance but lacking in dyeability. In the denim abrasion systems, it is used to lessen the incidence of cracks and streaks which occurs when chemicals and stonewashing is used. For commercial desizing, lipases are usually prepared in combination with α-amylases (Raja et al., 2012).

1.2.8.2 Lipases in Detergent Industry

Lipases are commercially important additives in detergents used in laundries and house dishwashers. According to Kurma et al. (2016b), about 32% of the total lipase sale is in the detergent industry where lipases are used as functional compounds in detergent formulation. The core features of lipases used in detergent formulation are solubility in water, stability under the conditions of washing (pH 10.0-11.0 and temperatures 30°C-60C), resistance to the other components of the formulation (surfactants and proteases) and broad substrate affinity. During washing, the lipase adsorbs on to the surface of fabric forming lipase-fabric complex which then acts on the oil stains and hydrolyses them. The enzyme reduces the load of chemical byproducts from detergents ditched into the environment. They enable lower temperature washing thereby saving energy. Lipases are biodegradable, leaving no harmful residues that can pose threat to microbial and aquatic life. Thus they are reliable environmental friendly alternative (Barros et al., 2010).

1.2.8.3 Lipases in Food and Beverage Industry

The major application of lipase in this industry is in dairy, baking, fruit juice, beer and wine production where they improve their physicochemical, organoleptic and nutritional properties. Lipases have been applied in the modification of nutritional content, flavour development, fragrance enhancement and improvement of texture of processed foods (Aravindan et al., 2007). Flavour modification is achieved by synthesis of esters of short chain fatty acids and alcohols through esterification and transesterification. The action of lipase on milk fats during cheese production  improves  its flavour, texture and  softness.  Lipolysed  milk fat  used  coatings  for chocolates, in margarine and butter improves the flavour and shelf life of these products (Jooyendeh  et  al.,  2009).  They  also  play a key  role in  fermentation  processes  involved  in manufacturing of sausage production. Lipases have been applied in refining of rice flavour, modification of soybean milk and improvement of aroma of wine as well as speed of its fermentation. Lipase from Candida rugosa finds application in ice cream, single-cell protein, carbohydrate esters and amino acid derivatives production (Aravindan et al., 2007). Moreover, Lipases have been used to inter-esterify tea seed oil which is then used to make chocolates. Tea seed oil being a byproduct of tea processing provides a cheaper alternative to cocoa butter used in marking chocolates thereby reducing the cost confectionaries. In addition lipases are used in biolipolysis – a process that involves removal of fats during processing of fish and meat, thereby ensuring a nutritionally healthier end product (Guerrand, 2017, Raveendran et al., 2018).

1.2.8.4 Lipases in Nutraceutical Industry

One of the major nutraceutical which contribute to healthy living in man is Polyunsaturated Fatty Acids (PUFAs). They are lipids with two or more double bonds on their fatty acid chains and play a wide range of physiological roles necessary for healthy living. The two major families of nutritionally important PUFAs are omega three (ω3) and omega six (ω6) fatty acids. These ω3 fatty acid (Eicosatrienoic acid, α-Linolenic acid, docosahexaenoic acid and Eicosapentaenoic acid), and ω6 fatty acid (linoleic and arachidonic acid) are in high demand as nutraceuticals. The demand is met by concentrating PUFAs in edible oil, alga, fish oil and byproducts of fish using lipases (Sangeetha et al., 2011).

1.2.8.5 Use of Lipases in Medical Field

Lipases are important enzyme markers and drug targets in the field of medicine. Elevated levels of lipase indicate certain infection and can be used as diagnostic tool.   For instance, acute pancreatitis and pancreatic injury can be determined by the level of lipases in blood. They are also applied in enzyme-linked serum triglycerides determination. Hence, they serve as biosensors in detecting triglycerides in food, pharmaceutical and blood samples, and in pesticide pollution analysis (Gurung et al., 2013).

Lipases from isolated Galleria mellonella were found by Annenkov et al. (2004) to have a bactericidal action on Mycobacterium tuberculosis. Lipases can be used as digestive aids in malabsorption disorders and as activators of tumor necrosis factor in treatment of cancers (Gurung et al., 2013).

1.2.8.6 Lipases in Pharmaceutical Industry

Lipase regiospecificity, enantioselectivity, and chemoselectivity enable them to be used in the removal of compounds and resolution of racemic mixtures in compounds of pharmaceutical importance. Lovastatin used in lowering of serum cholesterol is synthesized from lipase isolated from Candida rugosa. Also, the synthesis of Diltiazem chloride involve the use of optically pure (-) trans-methoxylphenyl glycidic acid methyl ester prepared from enantiospecific lipase from Serratia marcescens. The drug is a calcium channel inhibitor and acts as a vasodilator in the treatment of vascular disorders (Zhao et al., 2008). S. marescens lipase is also used for the racemic bioresolution of the non-steroidal anti-inflammatory drug ketoprofen (Sangeetha et al., 2011). Moreover, lipases show excellent stability in the presence of organic solvents, in which the pharmaceutical substrates are dissolved (Barros et al., 2010).

1.2.8.7 Lipases in Cosmetics and Fragrances

Skin care products use vitamin A and its derivatives as additives. These retinoids are commercially prepared from immobilized lipases. They are also used in synthetic hair products preparation (Gurung et al., 2013). A lot of high value products used as adhesives, lubricants and coatings in cosmetics and personal care industry are derived from fatty acids produced from hydrolysis of oil by lipases (Sangeetha et al., 2011).

Also, sweet smelling esters such as ethyl acetate, ethyl butyrate, ethyl methyl butyrate, ethyl valerate and ethyl caprylate used as fragrance are synthesized by esterification reactions catalysed by lipases (Ahmed et al., 2010).

1.2.8.8 Lipases in Biodegradation and Environmental Management

In the coastal environment, ecorestoration and enzymatic industrial waste oil processing has been achieved using microbial lipases to degrade oil spills. Amro and Soheir (2009) recommend lipases of Pseudomonas aeruginosa for castor oil degration. Lipases of Bacillus subtilis, Bacillus licheniformis, Bacillus amyloliquefaciens, Serratia marcescens, Pseudomonas aeruginosa, and Staphylococcus aureus were reported to degrade effluents from palm oil mill, diary, slaughter house, soap industry, and domestic waste water. (Prasad and Manjunath, 2011).

1.2.8.9 Lipases in Biodiesel Production

The application of lipase in biodiesel production has shown great prospects in quest for biodegradable, non-toxic and renewable energy source. It offers a host of advantages. The glycerol by-product glycerol could easily be removed without the requirement of a complex separation process. Moreover, free fatty acids used as the raw material are completely converted into alkyl esters eliminating the need of product purification. The choice of the best enzyme sources and optimization of the substrate and solvent concentrations, temperature, acyl migration and viscosity are important characteristics for lipases application in biodiesel production (Barros et al., 2010).

1.2.8.10 Lipases in Leather Industry

Soaking, bating and degreasing stages of leather processing employ lipases. Lipases alongside proteases are employed in the soaking stage of leather processing where they replace the use of soda ash or sodium tetrasulphide in the presence of surfactant. In addition, lipases and proteases are formulated in compatible surfactant for degreasing of leather. This is done to remove residual grease left after liming process. Degreasing breaks down protein membrane of fat sac, remove the fat and emulsify it in solvent (Parameswaran et al., 2013).

1.2.8.10 Lipases in Paper and Pulp Industry

Paper and pulp industry share 11% of the global industrial enzyme market (Parameswaran et al.,

2013). Environmental regulatory pressures have prompted the pulp and paper industry to adapt new technology to eliminate the presence of various contaminants in the bleaching plant effluents. Lipases are one the enzymes applied in this endeavour. The enzyme is more efficient in removal of pitch when compared to the use of chemical pulping or bleaching. Pitch is the sticky black resinous constituent of wood which deposit on pulp and pulping machineries thereby negatively impacting the pulping process and the quality of the pulp produced. Hydrolysis of triglycerides in wood resin to fatty acid and glycerol makes the resin less viscous, improves quality of pulp and reduces pitch deposition (Liu and Kokare 2017; Ramnath et al., 2017b).

1.3       Aspergillus niger

Aspergillus species refer to a genus of fungi with a global distribution. The subgenera are divided into anamorphic and telemorphic species (Suhaib et al., 2012). The colour of Aspergillus varies from species to species and has been used in physical identification of the organism. Aspergillus are generally consider safe, though some species of have been implicated in different fungal infections.  Among the important species of the organism are Aspergillus niger, A. fumigates and A. flavus (Rhodes, 2006).

Aspergillus niger is a common species of the genus Aspergillus. They are black in colour hence the name of the fungi. They are among the most common agents of food spoilage and biodegradation. Hence can they can be easily isolated from spoilt foods. They fungi generally regard as safe, have found extensive use in microbial biotechnology (Samson et al., 2007). The filamentous fungi are aerobes that have been reported to have adapted in environments of wide temperature (6-47°C) and pH ranges (2-10). These properties coupled to their profuse sporulation ensure their wide distribution. However, they grow better in warm and humid environments (Schuster et al., 2002).

1.3.1    Taxonomy of Aspergillus niger

Domain:          Eukaryota Kingdom:       Fungi Phylum:          Ascomycota

Subphylum:     Pezizomycotina

Class:              Eurotiomycetes Order:             Eurotiales Family:           Trichocomaceae Genus:            Aspergillus Species:          Aspergillus niger Source: (Samson et al., 2007)

1.3.2    Identification of Aspergillus Species

The common used methods of identification of Aspergillus species are based on morphological properties of the colony and microscopic examinations. These methods are relatively easier, cheaper and faster, and do not require complex instrumentation (McClenny, 2005). The major drawback is the absence of a way to differentiate among colonies that present similar morphological character especially when they are of similar species but different strains. To combat this drawback, molecular characterization have become increasing the method of choice in identification of species (Li et al., 2013).

1.3.2.1 Morphological Identification of Aspergillus species

Morphological features of Aspergillus culture have been studied. The key macroscopic features used in species identification are the colony diameter, colour, exudates and colony texture. Microscopic identification on the other hand use characteristics such as conidial heads, stipes, colour, length, vesicles shape and seriation, metula covering, conidia size, shape and roughness. These features are usually observed after seven days of culturing (Diba et al., 2007).

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