Overview of Fungal Lipase: A Review

Overview of Fungal Lipase: A Review

Abhishek Kumar Singh & Mausumi Mukhopadhyay

Received: 7 April 2011 /Accepted: 26 October 2011 /Published online: 10 November 2011# Springer Science+Business Media, LLC 2011

Abstract Lipases (triacylglycerolacyl hydrolases, EC3.1.1.3) are class of enzymes whichcatalyze the hydrolysis of long-chain triglycerides. In this review paper, an overviewregarding the fungal lipase production, purification, and application is discussed. Thereview describes various industrial applications of lipase in pulp and paper, food, detergent,and textile industries. Some important lipase-producing fungal genera include Aspergillus,Penicillium, Rhizopus, Candida, etc. Current fermentation process techniques such asbatch, fed-batch, and continuous mode of lipase production in submerged and solid-statefermentations are discussed in details. The purification of lipase by hydrophobic interactionchromatography is also discussed. The development of mathematical models applied tolipase production is discussed with special emphasis on lipase engineering.

Keyword Lipase . Enzyme . Modeling . Production . Purification

Historical Background

Lipases (triacylglycerol hydrolases, E.C. 3.1.1.3) are a class of hydrolase which catalyze thehydrolysis of triglycerides to glycerol and free fatty acids (Fig. 1). In addition, lipasescatalyze the hydrolysis and transesterification of other esters as well as the synthesis ofesters and exhibit enantioselective properties [1]. The ability of lipases to perform veryspecific chemical transformation (biotransformation) has made them increasingly popular inthe food, detergent, chemical, and pharmaceutical industries [2, 3]. Lipases have emergedas one of the leading biocatalyst/bio-accelerators with proven potential for contributing tothe multibillion dollar underexploited bioindustry, using both in situ lipid metabolism andex situ multifaceted industrial applications [4, 5]. Fungi able to produce lipases is found inseveral habitats, including soils contaminated with oils, wastes of vegetable oils, dairyproduct industries, seeds, and deteriorated food [6, 7]. Furthermore, the rapid developments

Appl Biochem Biotechnol (2012) 166:486–520DOI 10.1007/s12010-011-9444-3

A. K. Singh :M. Mukhopadhyay (*)Department of Chemical Engineering, Sardar Vallabhbhai National Institute of Technology,Surat 395007( Gujarat, Indiae-mail: [email protected]

of molecular biology techniques, as well as the availability of more reliable high-throughputscreening methods have improved the utility that lipase offer for organic synthesis.Currently, biocatalysts of second generation are being produced by adapting a wild-typeenzyme to a desired application [8–11]. The new strains of lipolytic microorganisms [12]are used. Some techniques have been developed to obtain higher conversions for highlyspecific enzymes for each application, improving the possibility of industrial applicationsfor lipase [3, 13].

Candida rugosa lipases have great significance for their diverse biotechnologicalpotential [5]. Existence of C. rugosa lipase isoforms has been reported by several authors[5, 14, 15]. Rhizopus species is mainly divided into three groups, Rhizopus oryzae,Rhizopus microsporus, and Rhizopus stolonifer. R. oryzae lipase, Rhizopus delemar lipase,and Rhizopus javanicus lipase have a substitution in the His134 and the Leu234 by an Asnand a Leu, respectively [16]. The lipase gene from R. stolonifer has been reported with 84%amino acid sequence identity to R. oryzae lipase. However, there is no report on themolecular characterization of lipase from R. microspores [17].

The LIP2 lipase from the Yarrowia lipolytica (YLLIP2) is an ideal candidate for enzymereplacement therapy due to its unique biochemical properties: It shows highest activity atlow pH values and is not repressed by bile salts. YLLIP2 belongs to the same gene familyas Thermomyces lanuginosus lipase, a well-known lipase with many applications in thefield of detergents and biotechnological processes [18]. Although these two lipases show ahigh sequence identity of 30.3%, they have relatively different biochemical properties.

The commercial uses of lipase comprise of a variety of different applications includingproduction of biopolymer and biodiesel, pharmaceuticals, agrochemicals, cosmetics andflavors, etc. [19]. Fungal lipases have gained special industrial attention due to theirstability, selectivity, and broad substrate specificity [20–22]. Lipases are ubiquitousenzymes with significant industrial potential since they have not only general advantagesof lipase, such as high catalytic activity, mild reaction conditions, environmentalfriendliness, and exquisite chemical, enantio- and regioselectivity. Some drawback becauselipases can work only under mild conditions is, i.e., lipase has much weaker activitycompared to the chemical catalysts, has high costs of production, and is time consuming.

Some of the major lipase-producing fungi are of the genera Mucor, Rhizopus,Geotrichum, Rhizomucor, Aspergillus, Humicola, Candida, and Penicillium [23–25]. Thethermophilic Mucor pusillus, Rhizopus homothallicus, and Aspergillus terreus are well-known as the producer of thermostable extracellular lipase. A lipase that is stable at highalkaline conditions and high temperature is rare; Mucor sp. produces an extracellular,inducible, alkalophilic, and thermostable lipase. There are few reports available so far withmolds with alkalophilic and thermostable lipase [26, 27].

The importance of lipases is observed by the number of published articles recently. Infact, over the last few years, there has been a progressive increase in the number of

Fig. 1 Hydrolysis of triglyceride by lipase

Appl Biochem Biotechnol (2012) 166:486–520 487

publications related to industrial applications of lipase catalyzed reactions performed incommon organic solvents, ionic liquids, or even in nonconventional solvents. There are lessreview articles on fungal lipases. This review is a compilation of reports on the fungallipase production, process operation, purification, and its industrial applications.

Applications

Fungal lipases constitute an important group of biotechnologically important enzymesbecause of the versatility of its properties and ease of mass production. Fungal lipases areversatile in its enzymatic properties and substrate specificity, which make it very attractivefor industrial applications. Lipases like most specialty and industrial enzymes areincreasingly produced via recombinant DNA technology [28]. Lipases are used in twodistinct fashions. It is used as biological catalyst to manufacture food ingredients and itsapplication as such in production of fine chemicals. Lipases are commonly used in theprocessing of fats and oils, food processing, leather, textile, detergents and degreasingformulations, paper manufacture, synthesis of fine chemicals, production of pharmaceut-icals, cosmetics [29], etc. Lipase can also be used to accelerate the degradation of fattywaste [30] and polyurethane [31]. Most of the industrial microbial lipases are derived fromfungi (Table 1).

In Textile Industry

The use of fungal lipase in textile industry is becoming increasingly important. Lipases areused to assist in the removal of size lubricants in order to provide the fabric betterabsorbency for enhanced levelness in dyeing. It also reduces the frequency of cracks andstreaks in the denim abrasion systems. Commercial preparations used for the desizing ofdenim and other cotton fabrics contain lipase enzymes [28].

In the textile industry, polyester has certain key advantages including softness, highstrength, washability, stain, stretch, machine abrasion, and is wrinkle resistance. Syntheticfibers have modified enzymatically for the use in the manufacture of yarns, fabrics, rugs,and textiles. It relates to modification of the characteristics of a polyester fiber so that suchpolyesters are additional susceptible to postmodification treatments [28].

In Detergent Industry

The most commercially important field of application for hydrolytic fungal lipases is asadditives in detergents in industrial laundry and household detergents [20]. Lipase canreduce the environmental load of detergent products as it save energy by enabling a lowerwash temperature to be used [32]. In 1994, Novo Nordisk introduced the first commerciallipase, Lipolase, which originated from the fungus T. lanuginosus and has been expressedin Aspergillus oryzae.

Fungal lipases function in the removal of stains from fabrics and are importantcomponents of detergent mixtures [33]. As early as 1988, the company Novo Nordiskdeveloped a fungal lipase, an enzyme capable of dissolving fatty stains. This enzyme isproduced naturally by a selected strain of Humicola, but at low concentration for industrialapplication. Traditional methods to enhance yield proved ineffective, so the gene coding forthis lipase has been cloned and inserted to A. oryzae. This fungus now produces the lipasein commercially significant yields so that it can be used in detergents, allowing better

488 Appl Biochem Biotechnol (2012) 166:486–520

washing performances and energy savings. Improved industrial lipase enzymes fights stains[28] more efficiently. Several fungus like A. oryzae [34], Candida sp. [35], R. oryzae [36],and Humicola lanuginosa are known to produce lipases under standardized conditionssuitable for detergent applications.

In Food Industry

The lipase used in food processing, industry is for the modification and breakdown ofbiomaterials. A large number of fat-clearing lipases are produced on an industrial scale.Most of the commercial lipases produced are utilized for flavor improvement for dairyproducts and processing of other foods, such as meat, vegetables, fruit, smoked carp, milkproduct, baked foods, and beer [37, 38].

Cocoa butter, a high value-fat (melting point of approximately 37 °C), contains palmiticand stearic acids. Quest-Lodrs Croklaan is commercialized the technology of synthesis ofsome of the less desirable fats in cocoa butter substitutes using an immobilized Rhizomucor

Table 1 Some commercially available lipases, their sources and industrial applications

Source Application Commercial name Producing company References

Humicolalanuginose

Detergent additive Lipolase TM Novo Nordisk [28]

C. cylindracea Food processing ChiroCLEC-CR Atlus Biologics [29, 32]

Lipase AY Amano

Lipase MY, Lipase OF-360 Meito Sangyo

Chirazyme® L-3 Boehringer Mannheim

Lipomod™ 34P-L034P Biocatalysts

C. rugosa Organic synthesis Lipase AY “Amano” 30 Amano [29]

Resinase® Novozymes

R. miehei Food processing Palatase® Novozymes [29]

T. lanuginosus Detergent additive Lipolase®, Lipolase® Ultra,Lipo Prime™, Lipex®

Novozymes [29]

A. niger Food processing Lipase A “Amano” 6 Amano [29]

Lypolyve AN Lyven

Rhizopus oryzea Food processing,oleochemistry

Lipase F-AP15 Amano [29]

Lipomod™ 627P-L627P Biocatalysts

R. niveus Oleochemistry Newlase F Amano [29]

M. miehei Food processing Piccnate Gist-Brocades [29]

Novo Nordisk

M. javanicus Food processing,oleochemistry

Lipase M “Amano” 10 Amano [29]

Penicilliumroquefortii

Food processing Lipomod™ 338P-L338P Biocatalysts [29]

Penicilliumcamemberti

Food processing,oleochemistry

Lipase G “Amano” 50 Amano [29]

Penicillium sp. Food processing Lipomod™ 621P-L621 Biocatalysts [29]

Candidaantarctica A/B

Organic synthesis Chirazyme®L-5 Boehringer Mannheim [32]

SP526 Nova-nordisk

Chirazyme®L-2 Boehringer Mannheim

SP 525 or Novozym 435b Nova-nordisk

G. candidum Oleochemistry Chirazyme®L-8 Boehringer Mannheim [32]

SP 524, Lipolase® Nova-nordisk


miehei lipase, which carries out a transesterification reaction replacing plamitic acid in palmoil with stearic acid to produce the desired stearic–oleic–stearic triglyceride [6, 39].Immobilized lipases are efficient, fast, accurate, and cost effective as sensors for thequantitative determination of triglycerol. Newlase, an immobilized lipase from Rhizpousniveus, specifically incorporates stearic acid at the sn-1 and sn-3 positions of triglycerides insunflower oil [29].

The Rohm and Haas Co. USA, which manufactures lipases, presumably lipase B, claimthat this lipase has used for improving the cheese flavor. Peters and Nelson [40] has studiedthat the addition of Candida lipolytica lipase enhanced the quality of blue cheese madefrom unhomogenized, pasteurized milk. The R. delemar lipase RH, produced by the TanabeSeiyaku Co., Japan has used lipases for enhancing flavors of dairy products, such as milk,cheese, butter, etc. For production of butter flavors, addition of lipase RH to fresh milk orcream at concentrations of 0.05–0.2% (w/w) followed by incubation at 40 °C for 2–5 h isrepeated. The reaction mixture is then heated to inactivate the lipase RH and spray-dried[40]. Some method is utilizes the immobilized R. miehei lipase for transesterificationreaction that replaces the palmitic acid in palm oil with stearic acid. Immobilized lipasesfrom Candida antarctica, Candida cylindracea AY30, H. lanuginosa, Pseudomonas sp.,and Geotrichum candidum are used for the esterification of functionalized phenols forproduction of lipophilic antioxidants in sunflower oil [41]. Lipases have generally used inthe production of a variety of products, ranging from fruit juices to vegetable fermentation[5]. Lipases facilitate the removal of fat from meat and fish products [6]. Cao et al. reporteda lipase-catalyzed solid-phase production of sugar fatty acid esters [42].

In Pulp and Paper Industry

A unique enzymatic pitch control system has been developed to degrade triglycerides present inmechanical pulp slurry [43]. This system has been used successfully at two paper mills of theJujo Paper Co. Japan, the Ishinomaki mill (1990) and the Yatusushiro mill (1991). At theYatusushiro mill (Ground wood pulp, 160 ton/day), the objective is to demonstrate the use ofenzymes with unseasoned wood logs. Resinase A from Novo Nordisk (liquid) obtained fromAspergillus sp. and lipase AYL (liquid) produced from C. rugosa and lipase powdersproduced from C. cylindracea are used in pulp and paper industry.

In Fat and Oleochemical Industry

The use of lipases in the oleochemical industry is enormous as it saves energy andminimizes thermal degradation during hydrolysis, esterification, acidolysis, alcoholysis,interesterification, and aminolysis (Fig. 2) [44, 45]. Although lipases are designed by naturefor the hydrolysis of the ester bonds in triglycerols in the presence of excess water, lipasescan catalyze the esterification reaction (ester synthesis) in a low-water environment.Hydrolysis and esterification can occur simultaneously in a process known as interester-ification. Depending on the substrates, lipases can catalyze alcoholysis (an acyl moietydisplaced between an acyl glycerol and an alcohol), acidolysis (an acyl moiety displacedbetween an acyl glycerol and a carboxylic acid), and transesterification (two acyl moietiesexchanged between two acylglycerols) [6]. Miyoshi Oil and Fat Co., Japan, reported thecommercial use of C. cylindracea lipase in the production of soap. Muco miehei (IM 20)and C. antarctica (SP 382) lipases have used for esterification of free fatty acids (FFAs) insolvent free or transesterification of fatty acid methyl esters in hexane with isopropylideneglycerols [46].


The esterification of oleic acid by ethanol using lipases from R. niveus and M.miehei as catalysts are evaluated in microaqueous, biphasic (n-hexane–water) andsurfactant-enriched biphasic systems containing different amounts of water [47]Immobilized M. miehei lipase in organic solvent catalyzed the interesterification reactionsfor production of vegetable oils such as corn oil, olive oil, sunflower oil, peanut oil, andsoybean oil containing omega-3 polyunsaturated fatty acids [48]. Freitas et al. hasreported that synthesis of monoglycerols in a medium solely composed of substrates byPenicillium camembertii lipase-catalyzed esterification of glycerol with fatty acids andabsence of solvent or surfactant [49].

Interesterification and hydrogenation techniques are useful in the preparation ofglyceride products for use in the production of margarine and butter. In the conventionalinteresterification reaction, interesterification is conducted in the presence of lipases fromRhytidiadelphus japonicus, Aspergillus niger, C. cylindracea, and G. candidum as catalystsare known; however, this process requires the presence of water to activate the lipase. Thepresence of water causes hydrolysis of interesterified glycerides with resultant decreases inyield of the glyceride product [50].

The current trend in the oleochemical industry is a movement away from using organicsolvents and emulsifiers on the different reactions involving alcoholysis, hydrolysis, andglycerolysis which have been carried out directly on mixed substrates using a range ofimmobilized lipases. This has resulted in high productivity as well as in the continuousrunning of the processes [51].

Fig. 2 Various lipase-mediated reactions


In Biodegradable Polymer Production

Lipases have recently become one of the most studied groups of enzymes. Lipases is usedas biocatalyst in the production of useful biodegradable compounds. 1-Butyl oleate issynthesed by direct esterification of butanol and oleic acid to reduce the viscosity ofbiodiesel in winter use. The mixture of 2-ethyl-1-hexyl esters is obtained in a good yield byenzymatic transesterification from rapeseed oil fatty acids for use as a solvent.Trimethylolpropane esters are also similarly synthesized as lubricants. Lipases can alsocatalyze ester syntheses and transesterification reactions in organic solvent systems hasopened up the possibility of enzyme catalyzed synthesis of biodegradable polyesters.Aromatic polyesters are produced by lipase biocatalysis [52].

In Pharmaceutical Industry

In the pharmaceutical industry, enzyme offers several advantages over chemical synthesis,thereby justifying the growing demands for lipase. These advantages include mild conditionsthat avoid isomerization, epimerization, racemisation, and rearrangement reactions; enantio-and regioselectivity; reuse of the immobilized lipase; overexpression of the lipase; economy ofthe process; andmutagenesis of the lipase for specific functions. The ability of lipases to resolveracemic mixtures by the synthesis of a single enantiomer is currently exploited for drugproduction by the pharmaceutical industry. Hirashima et al. [53] has carried out the hydrolysisof acyl bonds at the 1-position of 1, 2 diacylglycerophospholipids for the purification ofplasmalogens with R. delemar lipase. Berrobi et al. [54] have filed a patent for pharmaceuticalpreparations that contain hyaluronidase and thiomucase enzymes in addition to lipases for usein skin inflammations. Lipase is a component of a hair-waving preparation in which itpromotes penetration of the preparation [53]. Mushrooms with medicinal impact are used forpharmaceutical products [55]. Few essential mushrooms with pharmacological properties areAgaricus brasiliensis, Ganoderma lucidum, Lentinula edodes, Coriolus versicolor, Pleurotusostreatus, Grifola frondosa, Termitomyces, etc. [56, 57]. The important biologically activesubstances are found in the mushrooms such as immunosuppressive, antimicrobial, antiviral,nematicide, and hypocholesterolemic agents [58].

As Diagnostic Tools

Lipases can be used as diagnostic tools and its increasing levels indicate certain disease.These are important drug targets or marker enzymes in the medical sector. The lipaseslevels in blood serum are used as a diagnostic tool for detecting situation such as pancreaticinjury and acute pancreatitis [59]. Lipase activity/level determination is also important inthe diagnosis of heart ailments [60]. The mushrooms are used for the cancer therapies, dueto its high tolerance and compatibility with the chemotherapy and radiotherapy. The fruitbodies of mushroom and its extracts are efficiently used as economically feasible optiondue to faster growth of fruiting body [58]. The significant bioactive substances originate inmushroom such as lentinan, schizophyllan, and PSK; they are all used in cancerchemotherapy as component of the drug cocktails with impressive results [58].

In Cosmetics

Lipases have potential application in cosmetics of its activities in surfactants and in aromaproduction [61]. Mono- and diglycerols are produced by esterification of glycerols and are


used as a surfactant in cosmetics and perfume industries. Immobilized R. meihei lipase hasbeen used as a biocatalyst. Unichem International (Spain) has production of isopropylpalmitate, isopropyl myristate and 2-ethylhexylpalmitate for used as an emollient inpersonal care products such as skin and sun-tan creams and bath oils. The company claimsthat the use of the lipase in place of the conventional acid catalyst enhances product quality,with minimum downstream refining [28]. Croda Universal Ltd. has manufacture wax esters(esters of fatty acids and fatty alcohols) with C. cylindracea lipase in a batch bioreactor.According to the manufacturer, the overall production cost is slightly higher compared toconventional method but with enhanced quality of the final product [28].

In Tea Processing

The quality of black tea is dependent on the dehydration, mechanical breaking, andfermentation of the tea shoots. During manufacture of black tea, enzymatic breakdown ofmembrane lipids initiate the formation of volatile products with characteristic flavorproperties, which emphasize the significance of lipid in flavor improvement during teaprocessing. Lipase from R. miehei improved the level of polyunsaturated fatty acid byreduction in total lipid content. R. miehei increased the formation of desired flavourcompounds [62].

Black tea fermented with Dabaryomyces hansenii, results in accumulation of majorvitamins, such as A, B1, B2, B12, and C in sufficient quantities to fulfill the recommendeddietary allowance. It also results in decrease of caffeine and tannins in large amount.Moreover, the theophylline is accumulated as a result of fermentation imparts abronchodilatory effect to the tea [63]. Tea fermentation with D. hansenii enhanced itsnutritional and medicinal values. Lipases produced from R. miehei are used to enhance theflavour of tea during tea processing.

In Medical Applications

The primary enzyme for fat metabolism is lipase and its deficiency poses dire consequencesto health. Lipase, being an activator of the tumor necrosis factor, is used in the treatment ofmalignant tumors [64]. Other therapeutic applications of lipases, along with othercomponents, are found for treatment of dyspepsias, gastrointestinal disturbances, cutaneousmanifestations of digestive allergies, etc. [65]. The lipase level in blood serum is adiagnostic indicator for conditions such as acute pancreatitis and pancreatic injury [59, 66].Lipase activity determination is also significant in the diagnosis of heart ailments [67].Lipases may be immobilized onto pH/oxygen electrodes in combination with glucoseoxidase, and these function as lipid biosensors and used in triglycerides and bloodcholesterol determinations [68].

As Biosensors

A biosensor based on the enzyme-catalyzed dissolution of biodegradable polymer films hasbeen reported [69]. Three polymer–enzyme systems are studies for use in the sensor: adextran hydrogel, which is degraded by dextranase; a poly (ester amide), which is degradedby the proteolytic enzyme α-chymotrypsin; and poly (trimethylene) succinate, degraded bya lipase. Potential fields of application of such a sensor system are the detection of enzymeconcentrations and the construction of disposable enzyme-based immunosensors, which usethe polymerdegrading enzyme as an enzyme label [69]. Radiolabelled polynucleotide


probes are used widely for the detection of complementary nucleic acids by specifichybridization. Within the last few years, different methods have been developed usingenzyme-labeled probes to avoid unstable and hazardous isotopes. The use of unstablephosphatase and peroxidase conjugates is avoided due to the composition of thehybridization mixture and the high temperature. By screening different hydrolytic enzymesto fit the special demands, fungal lipases turned out to be the most practical [70].

Lipases are fast, efficient, and accurate as sensors for the quantitative determination oftriglycerol. The basic concept of using lipase as biosensors is to generate glycerol from thetriglycerol in the analytical sample and to quantify the released glycerol by an enzymaticmethod [28]. This principle enabled the physician to diagnose the patients withcardiovascular complaint. C. rugosa lipase biosensor from C. rugosa is developed as aDNA probe [71].

In Degreasing of Leather

Degreasing is an essential step in the manufacturing of glove and clothing leather. In thisprocess, there is removal of excess natural fats from greasy skins. The presence of naturalgrease in raw hides and skins, especially wooly sheep skins, results in various defects, viz.uneven dyeing and finishing, fatty spues, waxy patches in alum-tanned leathers, and pinkstain on wet blues [72] . Yeshoda et al. [73] is used as a lipase for the degreasing of woolysheep skins, pH range of 3.2–3.6 at 37 °C for 1 h. Yeshoda et al. [74] has also reported thatdegreasing and bating could be carried out simultaneously in the pH range of 7.8–8.0. Anacid lipase from Rhizopus nodosus has noticed to be very effective in the degreasing ofsheep skins [75].

In Waste Treatment

Lipases are utilized in activated sludge and other aerobic waste processes where thin layersof fats constantly removed from the surface of aerated tanks to permit oxygen transport.This skimmed fat-rich liquid is digested with lipases [76], such as that from C. rugosa.Lipases are also assisting the usual performance of anaerobic digesters [77]. Effectivebreakdown of solids and the clearing and prevention of fat blockage or filming in wastesystems are important in many industrial operations. Examples include: (1) sewagetreatment, cleaning of holding tanks, septic tanks, and grease traps; (2) degradation oforganic debris—a commercial mixture of lipase, cellulase, inorganic nutrients, wheat bran,etc. [78]. Jeganathan et al. [79] evaluate the hydrolysis of wastewater with high oil andgrease (O&G) concentration from a pet food industry using immobilized C. rugosa lipase(CRL) as a pretreatment step for anaerobic treatment through batch and continuous-flowexperiments. Preliminary anaerobic respirometric experiments confirmed the biodegrad-ability of wastewater pretreated by CRL and the O&G are reduction with and withoutpretreatment.

Leal et al. [80] has reported anaerobic treatment of synthetic model dairy wastewaterwith and without prehydrolysis by solid enzyme preparation from Penicillium restrictum. Itis found that the performance of anaerobic treatment of wastewater with and withoutprehydrolysis almost comparable when the concentration of oil and grease increase to600 mg L−1. However, when the concentration is 1,000 mg L−1, the wastewater treatmentcomplemented with enzymatic hydrolysis promoted a slight improvement on COD in thereactor fed with the hydrolyzed effluent. Margesin et al. [28] have found that monitoring ofsoil fungal lipase activity is a valuable indicator of diesel oil biodegradation in freshly


contaminated, fertilized, and unfertilized soils. Fungal species is used to degrade oil spillsin the coastal environment, which may improve ecorestoration as well as in the enzymaticoil processing in industries.

Rigo et al. [81] has studied a comparison between the noncommercial lipase preparationfrom P. restrictum produced in solid-state fermentation (SSF) and Lipolase 100T (immobilizedlipase from T. lanuginosus, Novozymes) in the hydrolysis of oil and grease in wastewater fromthe swine meat industry [81]. The optimum conditions of hydrolysis for every enzyme areobtained through statistical analysis methodology. The authors have proposed that thenoncommercial lipase preparation is more suitable for hydrolysis of meat industry effluentsdue to higher hydrolytic rates. Furthermore, its cost is low since it has crude enzymaticpreparation produced from agro-industrial residues [82]. Anonymous [83] used the lipase forthe degradation of wastewater contaminants such as olive oil from oil mills. Another importantapplication has been reported for the degradation of polyester waste, removal of biofilmdeposits from cooling water systems, and also purifying the waste gasses from factories.

In Biodiesel Production

In enzymatic transesterification, higher yields are achieved for biodiesel production withusing refined oil compared to crude oils. This is due to the presence of phospholipids in thenonrefined oil, which affect the interaction between lipase and substrate, since they possiblyocclude the pores of the support, in the case of using an immobilized lipase. Therefore, atleast the oil-degumming step is conducted before transesterification reactions in order toobtain a better production of biodiesel [84]. Kaieda et al. [85] has investigated themethanolysis of a soybean oil and methanol using R. oryzae lipase in a solvent-free reactionsystem (Fig. 3). It is anticipated that such soybean oil methyl esters is used as biodiesel fuel.The oil is first hydrolyzed to FFAs and partial glycerides and the FA produced is thenesterified with methanol. R. oryzae lipase is considered to exhibit 1(3)-regiospecificity, acertain amount of 1,3-diglyceride is obtained during the hydrolysis and methanolysis ofsoybean oil by R. oryzae lipase solution.

Matassoli et al. [86] has studied the enzymatic alcoholysis of crude palm oil with methanoland ethanol using immobilized lipases (Lipozyme RM IM, Lipozyme TL IM; 3% w/w) with amolar ratio of ethanol/oil of 3:1 at temperature of 50 °C. For the evaluation of the effect oftemperature on lipase-catalyzed biodiesel production, a semiempirical model has proposed byBrusamarelo et al. [87]. Brusamarelo et al. has investigated soybean biodiesel productionusing the commercial product Novozym 435 within the temperature range of 45–70 °C,observing the highest yield (92%) when 65 °C has been used. Pandey [88] has reported themethanolysis of a waste oil mixture (containing 1,980 ppm of water and 2.5% FFAs) usingimmobilized lipase from C. antarctica, and considering three steps of substrate addition.Soetaert and Vandamme [89] reported the use of the lipases from M. miehei and C. antarcticain the transesterification of various oils using hexane as solvent and found that the lipase from

Fig. 3 Biodiesel production by enzymatic transesterification


M. miehei is more efficient in converting primary alcohols (methanol, ethanol, propanol, and1-butanol) with yields between 95% and 98%, whereas lipase from C. antarctica is moreproper for the conversion of secondary alcohols (isopropanol and 2-butanol) with yieldsbetween 61% and 84%. In the absence of the solvent, the yields of methyl and ethyl esters aredecreased, particularly with methanol, with yield reduction up to 19%. Pandey [88] hasreported the use of some specific and nonspecific lipases (from C. rugosa, Pseudomonascepacia, and Pseudomonas fluorescens) in biodiesel production. The nonspecific lipasespromoted the highest yields of methyl esters when a molar ratio of 3:1 of methanol/oil isused. Specific lipases need gradual addition of methanol to achieve high yields (between 80%and 90%) and this is probably due to acyl migration of sn-2 to sn-1, which occursspontaneously in glycerides. Huang et al. has studied the methanolysis of soybean oil utilizingR. miehei displaying Pichia pastoris whole-cell biocatalyst. The optimal reaction conditionsare: temperature, 55 °C; water activity, 0.66; the ratio of isooctane to oil, 2:1 (v/v); and thewhole-cell biocatalyst, 40% (w/w, soybean oil) [90].

Sources

Lipases are ubiquitous in nature and are produced by several plants, mammals, andmicroorganisms. Lipases of microbial origin, like bacteria, fungal, and yeast, represent themost widely used class of enzymes in biotechnological applications and in organic chemistry. Alist of the common potential fungus for lipase production in fermentations is presented inTable 2 [91–107].

Table 2 Screening of various fungal strains for lipase production through fermentation

Microorganism Time (h) Lipaseactivity

Type offermentation

Raw material Reference

Penicillium aurantiogriseum 48 25 UmL−1 SmF Soybean oil [91]

R. rhizopodiformis 24 43.0 UmL−1 SSF Olive oil cake–Bagasse [92]

R. pusillus 25 10.8 UmL−1 SSF Olive oil cake–Bagasse [92]

P. restrictum 24 30 Ug−1 SSF Babassu oil cake [93]

P. simplicissimum 36 30 Ug−1 SSF Babassu oil cake [94]

Rhizopus oligosporus TUV-31 48 76.6 Ug−1 SSF Egg yolk [95]

Rhizopus oligosporusISUUV-16

48 81.2 Ug−1 SSF Almond meal [96]

Aspergillus carneu 96 12.7 UmL−1 SSF Sunflower oil [97]

C. cylindracea 179.5 23.7 UmL−1 SmF Oleic acid [98]

C. rugosa 50 3.8 UmL−1 SmF Olive oil [99]

P. verrucosum 48 40 Ug−1 SSF Soybean bran [100]

Geotrichum sp. 24 20 UmL−1 SmF Olive oil [101]

R. homothallicus 12 826 Ug−1 SSF Olive oil [102]

P. chrysogenum 168 46 UmL−1 SSF Wheat bran [103]

Fusarium solani FS1 120 0.45 Umg−1 SmF Sesame oil [104]

P. simplicissimum 48 21 Ug−1 SSF Soy cake [105]

Aspergillus awamori 96 495 UmL−1 SmF Rice bran oil [106]

C. cylindracea NRRLY-17506 175 20.4 UmL−1 SmF Olive mill wastewater [107]


Most fungal lipases are of considerable commercial importance for its bulk production.Although a number of lipase-producing fungi are recognized as follows: Rhizopus sp.,Penicillium sp., Aspergillus sp., Mucor sp., Ashbya sp., Geotrichum sp., Beauveria sp.,Humicola sp., Rhizomucor sp., Fusarium sp., Acremonium sp., Alternaria sp., Eurotriumsp., Ophiostoma sp., etc [108]. Lipase production by fungi varies according to the strain,the composition of the growth medium, cultivation conditions, temperature, pH, and thekind of carbon and nitrogen sources [109] used. In comparison to plant and mammalianlipases, microbial ones are most suitable for industrial applications due to its ease ofproduction, relatively inexpensive fermentation, and stability in organic solvents. Theindustry continues to look for economical new sources of lipases with varying catalyticcharacteristics which stimulates the isolation and selection of new strains [110]. Lipase-producing fungi can be isolated from different habitats such as industrial wastes, vegetableoil processing factories, dairy plants, soil contaminated with oil and oilseeds, compostheaps, coal tips, hot springs [6], etc. Isolation of 860 lipase-producing filamentous fungalstrains from soil samples from around the world using enrichment culture techniques havebeen reported [111]. An agar plate medium containing olive oil or tributyrin emulsionprepared in gum Arabic solution has been employed for isolation and growth of filamentousfungi in primary screening assay. Nine strains have been selected showing the best lipaseyields and the best reproducibility in liquid medium cultures.

Extracellular lipase production by isolated strain of Aspergillus sp. in soil samples fromdifferent regions of Turkey is studied by Cihangir and Sarikaya [109] with activity of 17 UmL−1. Colen et al. [112] has isolated 59 lipase-producing fungal strains from Braziliansavanna soil using enrichment culture techniques. Among these, the most productive strain,identified as Colletotrichum gloesporioides is produced 27,700 UL−1 of lipase. Isolation of20 lipase-producing fungal species from Ranchi, India is reported [113]. Three fungi A.niger KG-2, R. oryzae KG-5, and R. oryzae KG-10 has been found to be excellentproducers of lipase; R. oryzae KG-5 has shown maximum activity of 48.66 IU underoptimized conditions. An extracellular lipase Penicillium wortmanii [114] with lipaseactivity of 12.5 UmL−1, Penicillium cyclopium with lipase activity of 38 UmL−1 [115],Alternaria brassicicola with an activity of 3.2 UmL−1 [116], and A. niger NRRL3 with anactivity of 325 UmL−1 [117] has also been produced. Lin and Ko [118] has studied theeffect of different media concentration on extracellular lipase production of Antrodiacinnamomea.

The lipase production by P. restrictum with a lipase activity of 30.3 Ug−1 initial dryweight [93] and by Rhizopus oligosporous with lipases activity of 48.0 Ug−1 of substratehas been studied [119]. Sun and Xu [120] has optimized the nutritional parameters affectingproduction of whole-cell synthetic lipase by Rhizopus chinensis with synthetic activity of24,447 Ukg−1 of substrate. Studies of the production of acidic thermostable lipase byPenicillium simplicissimum strain grown in castor bean waste with maximum lipase activityof 44.8 Ug−1 [121], by Rhizomucor pusillus and Rhizopus rhizopodiformis with maximumproduction of 1.73 and 0.97 UmL−1, respectively, using sugar cane bagasse has beenobserved. The mixture of olive oil cake and sugar cane bagasse with maximum activity of43.04 and 10.83 UmL−1 has been obtained by R. rhizopodiformis and R. pusillus,respectively [92]. The production of lipase by A. niger strain MTCC 2594 using gingelly oilcake with a lipase activity of 363.6 Ug−1 [122] has also been reported.

A quantitative comparison between submerged fermentation (SmF) and SSF is difficultdue to the difference in the methods used for determining the lipase activity. However,some qualitative information presented in the literature can be of interest. In SmF, the lipaseproduction by Rhizopus arrhizus [123] with a maximum lipase activity of 315 UmL−1 is


reported. Studies of acidic lipase production by mutants of A. niger NCIM 1207 in SSFwith lipase activity of 630 IU g−1 is also reported [124]. Studies of acid and thermostablelipase production in SSF by a wild-type Brazilian strain of P. simplicissimum withproduction of about 90 Ug−1 in 72 h, with a specific activity of 4.5 Umg−1 of total proteins[125] has also been reported. Studies of extracellular lipases production by thermotolerantR. homothallicus with lipase activities of 10,700 Umg−1 (40 °C) and 8,600 Umg−1 (30 °C),by SSF and SmF, respectively, with the thermal stability of half-lives of 0.72 h in SSF and0.44 h in SmF [126] has been observed. The lipase activities of 12.1 and 17.4 Ug−1 in SmFand SSF, respectively, by cultivation of P. restrictum [127], A. niger J-1 with lipaseactivities of 4.8 and 1.46 IU mL−1, by SSF and SmF, respectively [128] and lipaseproduction by A. niger NCIM 1207 in both SmF and SSF with maximum lipase activity of630 IU g−1 dry solid substrate in SSF [124] has been reported.

Wolski et al. [129] has reported the use of response surface methodology to optimize thelipase production by SmF using immobilized cells of a newly isolated Penicillium sp. withlipase activity as high as 20.96 UmL−1 in 120 h of fermentation, higher than the activityobtained by the same free cells of this microorganism before immobilization. Theimmobilized cells of A. niger have used to produce lipase and similar activities for bothfree and immobilized cells cultivations with 4 UmL−1 has observed [24]. Yang et al. [123]has studied the repeated-batch lipase production by immobilized mycelium of R. arrhizus inSmF. The lipase productivity increased from 3.1 to 17.6 UmL−1 h−1, changing the processfrom batch to repeated-batch mode. The immobilized whole cell of R. arrhizus [130] onpolyurethane foam is used for lipase production with constant rate throughout severalrepeated batch experiments.

Isolation and Screening of Lipase Producing Fungi

Lipase-producing fungi has found in diverse habitats such as soil contaminated with oil, oilseeds, vegetable oil processing factories, decaying food, dairies, and industrial wastes[131]. The filamentous fungi are identified in-house by using mature cultures on suitablemedia (standard potato dextrose agar and oat meal agar) in order to ensure a gooddevelopment of taxonomically relevant features and following the identification keysprovided by von Arx [132], Domsch et al. [133], and specific methodology by Singler andCarmichael [134] in the case of Ovadendron sulphureo-ochraceum. Preliminary screeningof lipolytic filamentous fungi has been carried out on BYPO [133], agar platessupplemented with olive oil or tributyrin emulsion prepared in gum Arabic solution.Culture plates have been incubated at 28 °C and periodically examined for 4 days. Coloniesshowing clear zones around them have picked out and screened for lipase production inliquid medium [111]. A simple and reliable method for screening of lipase-producingfungal species has described by Kumar et al. Screening of lipase producers on agar plates,containing Bromophenol blue dye (pH 4.6) is frequently done by using olive oil as asubstrate and clear zones around the colonies indicate production of lipase. Colen et al.[112] have used potato dextrose agar plate to screen of fungal strains. Culture platesinoculated with the 59 fungal strains have been incubated at 30 °C for 72 h and the radiusof the colonies and the radius of the clear hydrolytic halos around them have beenmeasured. Toscano et al. [135] have used plates of tributyrin agar to screen for lipase-producing fungal species and mutant strains. Screening systems making use of olive millwastewaters substrates have also been described. Other versions of this method are alsoreported [136]. Screening for lipase-producing fungal species and mutant strains has carriedout on tributyrin agar on the basis of the coefficient K, which affects the ratio between the


diameter of the clear halo around the colony and the diameter of the respective colonymeasured on the seventh day at 30 °C. D'Annibale et al. [136] used the olive millwastewater (OMW) as substrates by its use as a possible growth medium for production oflipase. To this end, strains of G. candidum (NRRL Y-552 and Y-553), R. arrhizus(NRRL2286 and ISRIM 383), R. oryzae (NRRL 6431), A. oryzae (NRRL 1988 and 495),A. niger (NRRL 334), C. cylindracea (NRRL Y-17506), and P. citrinum (NRRL 1841 and3754, ISRIM 118) have been screened. All strains are able to grow on the undiluted OMWproducing extracellular lipase activity.

Lipase Assay

Lipases are known to hydrolyse triglycerides and give to FFA and glycerol. As with allreactions catalyzed by enzymes, activity measurements can be carried out using differentphysico–chemical methods by monitoring the disappearance of the substrate or by therelease of the product. Therefore, the assay methods are involved for measuring thehydrolytic activity as well as the detection of lipase. The methods are classified as: (1)titrimetry [137], (2) spectroscopy (photometry, fluorimetry, and infrared) [138], (3)chromatography [139], (4) interfacial tensiometry, (5) radio activity, (6) conductimetry,(7) turbidimetry, (8) immunochemistry [140], and (9) microscopy. Tributyrin plate assayand titrimetry are the most commonly used methods for screening of lipase producers andestimation of lipase activity [141, 142], respectively. The extensive reviews by Beisson etal. [143] and Gupta et al. [144] describe in detail the different methods available for lipaseassay. The most commonly used lipase assay protocol is the titrimetery assay using olive oilas a substrate because of its accuracy, simplicity, and reproducibility. A few spectropho-tometric assays are based on methods which render color to FA released after hydrolysis oftriglycerols [145].

Immobilization of Lipases

Lipases used in detergents and several other applications are not immobilized; however, anincreasing number of speciality applications of lipases in synthesis and biotransformationdemand an immobilized biocatalyst for efficiency of use. Immobilization is favored as itcan easily control the enzymatic process, purity of the products, and for its reusabilityfeature [146, 147]. Moreover, there is the possibility of lipase in an open form and capableof showing improved activity in immobilized condition [148]. Different techniques forimmobilization of lipases, such as physical adsorption, covalent bonding to a solid support[149], and physical entrapment within a polymer matrix support [150] have been used toprotect the lipase from the nonpolar solvent environment and enable its reuse. Among theimmobilization techniques, the adsorption is one of the simplest methods with a highercommercial potential than other methods. Although the immobilization of lipases has someadvantages, the selection of the support is a prominent factor influencing the enzymaticreactions [150]. G. candidum lipases A and B have been immobilized on Accurel EP 100porous polypropylene supports, precoated with ovalbumin to increase stability in organicsolvents and at elevated temperatures [151]. Bosley and Clayton [152] have employedhydrophobic controlled pore glasses to immobilize R. miehei lipase. Reetz et al. [153] haveused sol–gel entrapment in silica gel to immobilize different lipases. Xu et al. [41] haveused methyl acrylate divinyl benzene copolymer to immobilize C. cylindracea lipase. Theimmobilized lipase had improved resistance to thermal denaturation than the native enzyme


[154]. Reetz et al. [155] have studied an immobilization procedure using alkyl silaneprecursors of the type R Si (OCH3)3 and mixtures of R Si(OCH3)3 and Si(OCH3)4 toimmobilize C. antarctica lipase. Yong et al. [156] used the porous hydrophilic magneticmicrospheres to immobilized C. rugosa lipase. Lipases from C. rugosa, P. fluorescens, andC. antarctica B have been immobilized onto chitosan and glutaraldehyde-pretreatedchitosan powders.

C. rugosa lipase has been immobilized in mesoporous rod-like silica and vesicle-like (oronion-like) silica supports by physical adsorption. Dumitriu et al. [157] immobilized thelipase from C. antarctica B (CALB) on MCM-36 by physical adsorption. The acylation ofalcohols (1-butanol and 1-octanol) by vinyl esters (vinyl acetate and vinyl stearate) has beenused as a test reaction in order to evaluate the catalytic activity of the MCM-36immobilized lipase. Monduzzi et al. [158] investigated the physical and chemicaladsorption of Mucor javanicus lipase on SBA-15 mesoporous silica. Also, lipase fromCALB has been successfully entrapped in the cage-like pores of siliceous mesocellularfoam using a pressure-driven method [159]. Moreno et al. [160] have been covalentlyimmobilized C. rugosa lipase A and B on agarose and SiO2. Ghiaci et al. [161] haveinvestigated the usability of modified bentonite as a supporting material for immobilizationof C. rugosa lipase. Chiou and Wu [162] studied the immobilization of C. rugosa lipase tochitosan supports containing hydroxyl groups activated with carbodiimide. Ghamgui et al.[151] have used CaCO3 to immobilized R. oryzae by physical adsorption. Deng et al. [163]have used the polypropylene hollow fiber microfiltration membranes to immobilize lipasefrom C. rugosa by adsorption. A glycopolymer, poly (α-allyl glucoside) has grafted ontothe membrane surface by the plasma-induced polymerization of α-allyl glucoside toimprove the surface hydrophilicity and biocompatibility of polypropylene hollow fibermicrofiltration membrane. Knezevic et al. [164] have covalently immobilized C. rugosalipase on two epoxyactivated acrylic particulate polymers, namely Eupergit C and EupergitC 250L. Gupta et al. [165] have used the polyvinyl alcohol photomodified polysulfonemembranes to immobilized C. rugosa lipase. C. rugosa lipase has immobilized onpolyacrylonitrile nanofibrous membranes with the advantages of improved mechanicalstrength and high thermal resistance [166]. Lipase from C. rugosa has been noncovalentlyimmobilized on bentonite. Rodrigues et al. [167] have studied the immobilization andstabilization of the lipase from T. lanuginosus on glyoxyl agarose beads. Lipase from thefungus T. lanuginosus has been immobilized in mesoporous spherical particles produced bythe newly developed emulsion and solvent evaporation method. Organically modifiedsmectite nanonclays (ORKUN, ORSWy-2, and ORLAP) can act as efficient supports forthe immobilization of a lipase with numerous biocatalytic applications such the lipase Bfrom C. antarctica. Tzialla et al. [168] have reported the immobilized lipase B from C.antarctica on smectite group nanoclays (Laponite, SWy-2, and Kunipia). Yigitoglu andTemocin [169] used the poly (ethylene terephthalate) grafted acrylamide (PET-g-AAm)fiber to immobilized C. rugosa lipase.

Substrates

Fungal lipases are mostly extracellular and their production is greatly influenced bynutritional and physicochemical factors such as temperature, pH, nitrogen and carbonsources, and presence of lipids, inorganic salts, and agitation and dissolved oxygenconcentration. The major factor for the expression of lipase activity has always been studiedas the carbon source, since lipases are large inducible enzymes. These enzymes are


generally produced in the presence of a lipid such as oil or any other inducer, such as fattyacids, triglycerols, hydrolysable esters, bile salts, tweens, and glycerol [170] though fewauthors have produced good yields in the absence of fats and oils [6].

Organic and inorganic nitrogen sources and essential micronutrients in the medium areto be carefully considered for growth and production optimization [51]. These nutritionalrequirements for microbial growth are fulfilled by several alternative media as those basedon defined compounds (synthetic medium) like sugars, oils, and complex components suchas yeast extract, meat extract, malt extract, peptone, urea, ammonium sulfate, ammoniumnitrate, and also agro-industrial residues containing all the components necessary for fungaldevelopment. The studies available in the literature covering this area are presented belowconsidering the kind of medium used.

Synthetic Medium

Increasing lipase production during the fermentation process is also an important step inindustrial application of these enzymes. Various methods are used to optimize thefermentation process to improve production of lipase. However, few efforts have beenmade to improve the fermentation process by searching the variation of mediumcomponents and internal interactions during the cultivation process.

Kaushik et al. [97] have used the response surface approach to study the production ofan extracellular lipase from Aspergillus carneus. Interactions are evaluated for five differentvariables like sunflower oil, glucose, peptone, agitation rate, and incubation period and 1.8-fold increase in production, with the final yield of 12.7 IU mL−1 reported. The influence ofdifferent culture conditions, like temperature, pH, carbon, nitrogen, mineral sources, andvitamins factors etc. on the production of lipase by A. cinnamomea [171] in submergedcultures has been investigated. Nine carbon sources, 14 nitrogen sources, 6 mineral sources,and 5 vitamins are varied. The authors have showed that 5% glycerol, 0.5% sodium nitrate,and 0.1% thiamine provided the best results. The lipase production reached 54 UmL−1 afterincubation for 17 days. Response surface methodology is also used to optimize the culturemedium for lipase production with Candida sp. 99–125 [172]. The lipase fermentation in a5-L vessel is reported as 9,600 IU mL−1. Martinez Ruiz et al. [173] reported the productionof lipase from Rhizopus sp. using perlite (as inert support) supplemented with urea, lactose,olive oil, etc. with lipase activity of 75 IU g−1 after 20 h.

Agro Industrial Residues

The lipase production by P. simplicissimum using soybean meal as substrate supplementedwith low-cost supplements like soybean oil, wastewater from a slaughterhouse (rich on oiland fat) corn steep liquor, and yeast hydrolysate [174] are investigated. Cultivation conditionsare optimized for the production of lipase by factorial design and response surfacemethodology. Soybean meal without supplements appears to be the best medium of thosetested for lipase production with lipase activity of 30 Ug per dry substrate (ds−1) has beenreported. Damaso et al. [175] has investigated the lipase production from A. niger mutant11T53A14 by SSF using agro-industrial residue supplemented with by-products from corn oilrefining process or olive oil. The best results are reported with wheat bran as a nutrient sourceand soapstock, stearin, and fatty acids as a lipase production inducer. The highest lipaseactivities using soap stock, stearin, and fatty acids are 62.7, 37.7, and 4.1 Ug ds−1,respectively. The factorial design and response surface analysis has studied to optimize lipaseproduction from enicillium verrucosum by SSF [100] using soybean bran as substrate and


lipase activity as high as 40 Ug−1 of dry bran has been reported. Mala et al. [176] havedeveloped an SSF for lipase production by A. niger MTCC2594 using wheat bran andgingelly oil cake as substrates and supplement, respectively, and the results showed thataddition of gingelly oil cake to wheat bran increased the lipase activity by 36% and theactivity is 384 Ug ds−1. The digital image processing technique [21] has been used to monitorthe biomass growth of A. niger in SSF for lipase production. The strain of A. niger iscultivated in SSF using wheat bran as support, enriched with 0.91% of ammonium sulfate.The addition of several vegetable oils (castor, soybean, olive, corn, and palm oil) has beeninvestigated to enhance lipase production. The columns were inoculated with a suspension of107 spores/g of substrate, incubated in a thermostatic bath at 32 °C, and aerated with saturatedair at a rate of 4 Lh−1. The experiment lasted 96 h and was monitored every 24 h fordetermining the glycosamine content and the lipase activity. A maximum lipaseactivity is obtained using 2% of castor oil. The immobilized esterification lipaseactivity was determined for the butyl oleate synthesis, with and without 50% v/vhexane, resulting in 650 and 120 Ug−1, respectively. The lipase was used fortransesterification of soybean oil and ethanol with maximum yield of 2.4%, after30 min of reaction. A combined substrate of wheat flour with wheat bran supported bothgood biomass and enzyme production by R. chinensis in SSF is reported [120]. Mahadiket al. [124] has demonstrated that several agro-industrial residues such as cassavabagasse, apple pulp, and beans residues supplemented or not can be used as substrates forlipase production by solid state fermentation with several microorganisms.

Production Processes

Fermentative processes have operated in batch, fed-batch, and continuous reactor. Themode of operation is, to a large extent, dictated by the characteristics of the product ofinterest. This section is considering recent applications of batch, fed-batch, and continuousprocesses to lipase production in SmF and SSF.

Batch Processes

Suitability of OMW to serve as a medium for lipase production by C. cylindracea NRRLY-17506 in batch cultures has been investigated. The maximum lipase production of 21.6 UmL−1 depending on the dissolved oxygen concentration in the medium and activity of20.4 UmL−1 at pH 6.5 is reported [107]. The highest lipase activity of 9.23 IU mL−1 hasbeen reported [136]. The kinetics of the synthesis of lipase by C. rugosa in a batchbioreactor has been reported [177]. The investigators have illustrated the influence of gas–liquid mass transfer coefficient on the cell growth and lipase productivity. To maintainsufficient oxygen concentration for the optimum cell growth and lipase activity, reactionhas carried out using triple impeller bioreactor at an operating speed of 600 rpm and atdifferent aeration rates. Gas flow rate of 50.34 cm3s−1 has yielded optimum production oflipase and the lipase activity enhanced 2.5-fold. The lipase productivity of G. candidum instirred tank and in airlift bioreactors was compared [178]. In the stirred reactor, theoptimum conditions of agitation and aeration for lipase production was maintained at300 rpm and 1 vvm, leading to an activity of 20 UmL−1 in 54 h of reaction and productivityof 0.39 UmL−1 h−1. For the airlift bioreactor, the best aeration condition has maintained as2.5 vvm, and lipase activity of 20 UmL−1 after 30 h of reaction and productivity is reportedas 0.64 UmL−1 h−1. In the absence of mechanical agitation, lipase yields of 20 UmL−1 has


been achieved in less time, resulting in about 60% greater productivity compared to thatobtained in the stirred reactor.

The lipase production by Y. lipolytica [179] in a stirred tank reactor at different agitationspeeds and air flow rates has been investigated. The most pronounced effect of oxygen onlipase production is determined by stirring speed. A maximum lipase activity has beendetected in the stationary phase at 200 rpm and an air flow rate of 0.8–1.7 vvm, when thelipid source has been fully consumed. Higher stirring speeds resulted in mechanical oroxidative stress, while lower speeds seemed to limit oxygen levels. An increase in theavailability of oxygen at higher air flow rates has increased lipid uptake and anticipation ofenzyme release in culture medium. The maximum lipase production has reported at200 rpm and 1 dm3 min−1.

The fungal lipase production by submerged fermentation has reported the lipaseproduction in conical flasks using a few grams of substrate [93, 100, 176, 180]. SSF hasoffered improved yield and product spectra compared to SmF. Compared to SmF, wherethere are variations in bioreactor configuration and improved yield and product spectra, SSFis mostly restricted to a packed-bed configuration. There is limited literature available onthe use of an intermittently agitated, rotating drum, and fluidized bioreactor for batchreactors. Cavalcanti et al. [180] has studied a 30-g fixed-bed bioreactor to improveproductivity and scaling-up of lipase production using P. simplicissimum in SSF. Theinfluence of temperature and air flow rate on lipase production was assessed by statisticalexperimental design, and an empirical model was proposed to validate the experimentaldata. Higher lipase activities have achieved at lower temperature and higher air flow rates.A maximum lipase activity of 26.4 Ug−1 was reported at 27 °C at an air flow rate of 0.8 Lmin−1. Mala et al. [176] has studied the lipase production using 100 g and 1 kg of A. nigerMTCC 2594 in batch fermentation resulting in enzyme activities of 95% in 72 h.

Repeated-Batch Processes

The repeated-batch processes combine the advantages of fed-batch and batch processes toconduct the process by long periods and improves the productivity compared to the batchprocess. Elibol and Ozer [130] used repeated fed-batch strategy to produce lipase fromimmobilized R. arrhizus cells. The maximum lipolytic activity was under 100 UmL−1

reported at 1 gL−1 glucose concentration. The addition of 0.5 gL−1 corn oil to thefermentation medium resulted in 2.5-fold higher lipase production compared to the controlwhere no oil was added. The storage stability of the enzyme has investigated and enzymeactivity reduction was 26% within 160 h. Yang et al. [123] studied the repeated-batchfermentations for lipase production by immobilized mycelium from R. arrhizus insubmerged fermentation. The times to replace the volume, the volume of the replacedmedium, and the optimal composition of the medium for replacing during the repeatedbatch fermentation have been optimized. Nine repeated batches were carried out in flasksfor 140 h and six repeated batches in a 5-L fermentor. The lipase productivity increasedfrom 3.1 UmL−1 h−1 in batch fermentation to 17.6 UmL−1 h−1 in repeated-batchfermentation, which has 5.6 times as high as that in batch fermentation. Benjamin andPandey [181] carried out experiments in batch and repeated-batch (fed-batch type) forlipase production using immobilized C. rugosa cells in packed-bed bioreactor (PBR). Amaximum enzyme activity (17.9 UmL−1) was obtained when the fermentation was carriedout in a PBR in repeated-batch mode using a feed medium containing Arabic gum andcaprylic acid, keeping the flow rate of the feed at 0.4 mL min−1 and allowing each cycle torun for 12 h.


Fed Batch Processes

In the fed batch process, nothing is removed from the reactor during the process, but one ormore nutrients are added in order to control the reaction rate by its concentration. The fed-batch processes are used to minimize the effects of the cell metabolism control and, mainlyto prevent the inhibition by substrate or metabolic products. Bioreactors scale up from 5 to800 L for the high-cell density fermentation of C. rugosa lipase in the constitutive P.pastoris expression system was studied [182]. The fermentation conditions for both lab andpilot scale were optimized. The exponential feed input combined with pH control issuccessful in small-scale reactor. In two-stage pilot-scale fermentation, the culturetemperature and pH, was considered effective. A lipase activity of approximately14,000 UmL−1 was reported in 48 h.

Different fed-batch cultivation strategies in a bioreactor for the production of lipase byR. oryzae from P. pastoris were compared by Surribas et al. [183]. Several drawbacks wasreported using a methanol nonlimited fed-batch. Oxygen limitation appeared at early celldry weight and high cell death was observed. Temperature-limited fed-batch model wasproposed to solve both problems. However, a methanol nonlimited fed-batch has resulted inbetter productivity. A culture medium with low concentration of salt was used to overcomecell death problems. A temperature-limited fed-batch was applied thereafter to solve oxygentransfer limitations. This combined strategy has resulted in lower productivities whencompared to a methanol nonlimited fed-batch. However, the cultivation has extended for alonger time and 1.3-fold purer final product was obtained mainly due to cell deathreduction.

Kim and Hou [98] cultivated C. cylindracea NRRLY-17506 to produce extra cellularlipase from oleic acid as a carbon source. Fed-batch cultures were carried out in a bioreactorto improve cell concentration and lipase activity compared to batch system. For theintermittent feeding, the final cell concentration and the lipase activity has 52 gL−1 and6.3 UmL−1 after 138.5 h of fermentation. Stepwise feeding was carried out to simulate anexponential feeding and to investigate the effects of specific growth rate on cell growth andlipase production. The highest final cell concentration was obtained as 90 gL−1 when theset point of specific growth rate is 0.02 h−1 with the highest lipase activity of 23.7 UmL−1.High specific growth rate has decreased lipase production in the latter part of fed-batchcultures, due to build-up of excess oleic acid. A fed-batch fermentation process wasdeveloped [184] to enable the production of large quantities of recombinant humanlysosomal acid lipase in Schizosaccharomyces pome. A feedback fed-batch system wasused to determine the optimal feed rate of 50% glucose solution as carbon source. At thetime of the initial consumption of glucose in the batch-phase culture, the nutrient supply hasautomatically initiated by means of monitoring the respiratory rate change. The obtainedprofile of the feed rate was applied to the feed-forward control fermentation. Finally, thecells have grown up to 50 gL−1 dry cell weight with the lipase expression of approximately16,000 UL−1.

The lipase production by C. rugosa in a bioreactor with different fed-batch operationalstrategies (1) constant substrate feeding rate and (2) specific growth rate control [185] wasinvestigated to optimize lipase activity. A constant substrate feeding rate strategy hasreported that maximum lipase activity of 55 UmL−1 reached at low substrate feeding rates,where as lipase tends to accumulate inside the cell at higher rates of substrate addition. Inthe second fed-batch strategy investigated, a feedback control strategy was developed basedon the estimation of state variables from the measurement of indirect variables, such ascarbon dioxide evolution rate by mass spectrometry. An on–off controller has been used to


maintain the specific growth rate at the desired value, adjusting the substrate feeding rate. Aconstant specific growth rate strategy afforded higher final lipase activity of 117 UmL−1 atlow specific growth rates. With a controlled specific growth rate, lipase production by C.rugosa has increased totenfold compared to a batch operation.

Continuous Processes

The lipase production using SSF and SmF thermotolerant R. homothallicus in a 50-gpacked-bed bioreactor at 40 °C and 50 mL min−1 of air flow rate was [126] reported. Amaximum lipase activity of 1,500 Ug ds−1 is achieved after 12 h of fermentation.D'Annibale et al. [137] has studied the lipase production by C. cylindracea NRRLY-17506using OMW in a bubble column reactor (3 L). For the suitability of OMW as a lipaseproduction medium, four different OMW samples are used. The effect of nitrogen wasinvestigated by adding to OMW-4 the same amount of nitrogen (0.63 gL−1) present as oneof the following compounds: NH4Cl, (NH4)2 SO4, NaNO3, and urea. The influence of oiladdition was evaluated by adding to OMW-4 one of the following oils (3.0 gL−1): olive oil,corn oil, and soybean oil. The lipase production of 430 UL−1 with lipase activity of9.23 IU mL−1 was reported.

The production of extra-and intracellular lipases in continuous cultures of C. rugosausing sole or carbon source mixtures and different C/N ratio [186] has been studied. Lipaseproductivity in continuous cultures has increased by 50% compared to batch fermentationand dependent on the dilution rate applied. The authors found that during nitrogenlimitation, lipase activity was suppressed. The production of extracellular lipase by theThermomycesl anuginosus in chemostat cultures at a dilution rate of 0.08 h−1 in relation todifferent ammonium concentrations in the feed medium is investigated [187]. Under steady-state conditions, three growth regimes have recognized and the production of severallipases from T. lanuginosus under nitrogen to carbon limitations has recorded.

Purification

Many fungal lipases have extensively purified and characterized in terms of their activityand stability profiles relative to pH, temperature, and effects of metal ions and chelatingagents. Most commercial applications of lipase require homogeneous lipase preparationswith a certain degree of purity to enable efficiency. Besides, purification of enzymes allowsdetermination of primary amino acid sequence and 3-D structure [188–190]. Further,purified lipase preparations are needed for biocatalytic production of pharmaceuticals,cosmetics, leather, detergents, foods, perfumery, medical diagnostics, and other organicsynthetic materials.

The main constraints in traditional purification strategies include low yields and longtime periods. Alternative new technologies such as membrane processes, immunepurification, and aqueous two-phase systems are gradually coming to the purification oflipases. The purification strategies employed are inexpensive, rapid, high-yielding, andamenable to large-scale operations for industry purpose. It has potential for continuousproduct recovery, with a relatively high capacity and selectivity for the desired product.Various purification protocols used for lipases have been reviewed many times [190],highlighting clearly the importance of designing optimal purification strategies for variousmicrobial lipases. The extent of purification varies with the order of the purification steps;and this aspect was evaluated through the different purification procedures used by various


investigators. In this article, up-to-date literature on purification of fungal lipases wassummarized.

Prepurification steps involve concentration of the cell-free culture broth containing thelipase enzyme by ultrafiltration, ammonium sulfate precipitation, or extraction with organicsolvents. Precipitation was used as a crude separation step, often during the early stages of apurification method, and gives a high average yield [189] and such enzyme preparations areused in detergent formulations. Increase in lipase activity depends on the concentration ofammonium sulfate solution used [191].

Purification methods used depends on nonspecific techniques such as precipitation,hydrophobic interaction chromatography, gel filtration, and ion exchange chromatography.Precipitation was used as a fairly crude separation step, often during the early stages of apurification method, followed by chromatographic separation. Affinity chromatographywas used in some cases to reduce the number of individual purification steps needed [192].Activity of lipase depends on the concentration of ammonium sulfate solution used [191].Precipitation method often gives average yield of 87% with limited purification and suchenzyme preparations are used in detergent formulations. However, for certain applications,such as synthetic reactions in pharmaceutical, food, and leather industry, furtherpurifications are needed. Since lipases are known to be hydrophobic in nature, with largehydrophobic surfaces around the active site, the purification of lipases can best be achievedby opting for affinity chromatography such as hydrophobic interaction chromatography.Affinity methods can be applied at an early stage but as the hydrophobic matrices areexpensive, alternatively ion exchange and gel filtration are usually preferred after theprecipitation step [193]. Although gel filtration has a lower capacity for loaded protein, itcan be used at an early stage in the purification or as one of the last steps for fine polishingof the protocol.

The extracellular lipase from Pythium ultimum was purified by ammonium sulfateprecipitation, by diethylaminoethyl-Sepharose CL-6B, and by Sephacryl S-200 chroma-tography [194]. The purified lipase with a molecular mass of 68,000 gmol−1 by sodiumdodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) was reported. However,chromatography under nondenaturing conditions has indicated that the molecular weight ofthe lipase as 270,000 gmol−1, suggesting that the enzyme may be a tetramer. Thepurification and enzymatic properties of three kinds of lipases, A, B, and C, produced by R.delemar are studied [195]. The concentrated lipase was then purified on Ultrogel andSephadex G-150 columns. Lipase Awas purified 106-fold with 11% recovery; lipase B waspurified 71-fold with 3% recovery, and lipase C was purified 47-fold with 4% recovery. Theenzymes have been found homogeneous as judged by electrophoresis with molecularweights of 76,000, 60,000, and 45,000 gmol−1 for the A, B, and C lipases, respectively.

Ohnishi et al. [196] has reported an A. oryzae strain that produced at least two kinds ofextracellular lipolytic enzymes, L1 and L2. The lipolytic enzyme L1 has purified fromculture filtrate to homogeneity by ammonium sulfate and acetone fractionation, ionexchange chromatography, and gel filtration. Lipase L1 has a monomeric protein (24,000 gmol−1 molecular weight) and preferentially cleaved all the ester bonds of triolein.Purification and enzymatic properties of two kinds of lipases, I and II, produced by R.niveus was studied by Kohno et al. [197] are using column chromatography on DEAE-Toyopearl. Lipase I consisted of two polypeptide chains, a small peptide with sugar moiety(A-chain) and a large peptide of 34,000 gmol−1 molecular weight (B-chain). The molecularweight of lipase II was estimated to be 30,000 gmol−1 and a single polypeptide chain.Suzuki et al. [198] used chromatography on hydroxylapatite, octyl-Sepharose, andSephacryl S-200 to purify lipase from R. japonicas NR 400. The enzyme has an apparent


molecular weight of 30,000 gmol−1 by SDS-PAGE and activity is 31%. The lipase obtainedfrom R. delemar was purified 10.3-fold with 30% recovery [199] by oleic acid affinitychromatography, CM-Sephadex. An extracellular lipase from R. oryzae was purified160-fold with 64% recovery [200] by acetone precipitation (80%), Sephadex G-100, and1,260-fold with 22% recovery [201] by ammonium sulfate fractionation, sulfopropyl-Sepharose, Sephadex G-75 and again on sulfopropyl-Sepharose. The lipase obtained fromR. arrhizus was purified 720-fold with 42% recovery [202] by ammonium sulfatefractionation and Sephadex G-100 gel filtration. The purified enzyme has an apparent molecularmass of 67,000 gmol−1 on SDS-PAGE. The lipase obtained from R. chinensis has been purifiedby CM-Cellulofine C-500, ether Toyopearl 650 M, Super Q Toyopearl and CM CellulofineC-500. The enzyme has an apparent molecular mass of 28,400 gmol−1 and recovery in activityof 27.6% [203]. An extracellular lipase from R. hiemalis was purified 2,200-fold usingultrafiltration, ammonium sulfate fractionation, Sephadex G-75, Q-Sepharose chromatography,and Sephacryl S-200 chromatography [186]. The enzyme has molecular weight of 49,000 gmol−1 by Sephadex G-75 and SDS-PAGE. Wu et al. [204] has purified lipase of R. miehei42-fold with 32% activity with molecular weight of 31,600 gmol−1 by ammonium sulfateprecipitation, phenyl Sepharose fast-flow hydrophobic interaction chromatography, and DEAE-Sepharose fast-flow anion exchange chromatography.

An extracellular lipase produced by a selected strain of M. miehei was partially purifiedin two forms, A and B, by Huge-Jensen et al. [205]. Lipase Awas purified 15-fold from thecrude powder with 57% recovery by DE-cellulose 52 chromatography and affinitychromatography on Con A–Sepharose column. Lipase B was purified from the crudepowder by DEAE-Sepharose and phenyl-Sepharose chromatography, with an eightfoldpurification and a recovery in activity of 32%. A triacylglycerol lipase from the conidia ofNeurospora crassa was purified by Sephadex G-100 column chromatography [206].Homogeneity was checked by PAGE. The enzyme with single band of 27,000 gmol−1 hassuggested the presence of two identical subunits. An extracellular lipase produced byNeurospora spp. TT-241 was purified 370-fold with an overall yield of 16% [207]. Thepurification steps included ammonium sulfate precipitation, G-100 gel filtration, andchromatography using Toyopearl phenyl-650 M and Ultrogel-HA. The apparent molecularweight by SDS-PAGE was reported at 55,000 gmol−1 at 30 °C. Sugihara et al. [208] haspurified an extracellular lipase A. niger by precipitation with ammonium sulfate,hydrophobic interaction (butyl Toyopearl 650 M), gel filtration (Sephadex G-75), anionexchange chromatography (DEAE-Sepharose CL-6B), and adsorption on hydroxyapatite.In this procedure, the lipase was purified approximately 600-fold over the crude extractwith an activity yield of 34%. The apparent molecular mass by SDS-PAGE and SephadexG-100 chromatography was reported to be 35,000 gmol−1. An acid-resistant lipase from A.niger has purified from a crude commercial preparation by a procedure including sizeexclusion on Bio-Gel P-100 and ion exchange on Mono Q [209]. The overall purificationhas improved 40-fold with 36% yield. The most effective step in the purification procedurewas Mono Q ion exchange chromatography which increased 5.6-fold in specific activity.An extracellular lipase from A. niger F044 was purified to homogeneity using ammoniumsulfate precipitation, dialysis, DEAE-Sepharose Fast Flow anion exchange chromatography,and Sephadex G-75 gel filtration chromatography. The overall purification was 73.71 with34% yield and molecular weight of 35,000–40,000 g mol−1 using SDS-PAGE. The pH andtemperature optimum for lipolytic activity of the lipase was 7 °C and 45 °C [193]. Thelipase enzyme obtained from Aspergillus sp. culture medium was partially purified bySephacryl S-100 gel filtration column. Purity of the enzyme was checked by SDS-PAGEand molecular weight determined as 88,000 gmol−1. It has detected that the enzyme


retained its stability between 30 °C and 70 °C [110]. A lipase of A. oryzae was purified byammonium sulfate fractionation, anion exchange chromatography using Econo-Pac Q,hydrophobic interaction, and two successive ion exchange chromatography steps employ-ing Mono Q. The purified enzyme has a monomeric protein with a molecular mass of 41 gmol−1 estimated by SDS-PAGE and 39,000 gmol−1 by gel filtration [210]. Similarly, Toidaet al. [211] purified A. oryzae lipase by ammonium sulfate fractionation, acetoneprecipitation, and anion exchange chromatography and gel filtration with 11% yield. Theenzyme has reported a glycoprotein with molecular weight of 25,000 gmol−1.

A lipase produced by A. terreus was purified to electrophoretic purity by means ofammonium sulfate and acetone precipitation, gel filtration (G-100) and ion exchangechromatography (Q-Sepharose). The molecular mass by SDS-PAGE was reported as41,000 gmol−1 with the specific activity of 250 Umg−1 of protein [212]. A lipase producedby A. carneus was purified by a simple two-step procedure involving ammonium sulfateprecipitation and hydrophobic interaction chromatography on octyl-Sepharose. The purifiedlipase has a molecular weight of 27,000 gmol−1. A lipase monomer of apparent molecularweight of 29,000 gmol−1 was purified to homogeneity from extracellular culture of A.nidulans WG312 by phenyl-Sepharose chromatography and affinity binding on linolenicacid agarose [213].

A lipase from P. camembertii U-150 was purified into four active fractions by a procedureinvolving ammonium sulfate fractionation, ethanol precipitation and aminooctyl-Sepharose,hydroxyapatite, and Con A–Sepharose column chromatographies [214]. However, a novelenzyme hydrolysing mono- and diglycerols was found in the culture filtrate of P. camembertiiU-150 by Yamaguchi and Mase [215]. The enzyme was separated into two forms (A and Benzyme) with almost the same molecular weight (37,000–39,000 gmol−1), amino acidcomposition and identical N-terminal amino acid sequence. B enzyme, a major component,was purified approximately 210-fold with an activity yield of 2.6%. The extraction and back-extraction of a lipase from crude extract of P. citrinum using AOT surfactant reversed micellesin isooctane was well-documented [216]. The effect of pH, ionic strength, and AOTconcentration on the protein forward and backward transfer at 20 °C was reported. Maximumprotein forward extraction (32%) was achieved at pH 4.0 with a 50 mmol L−1 acetate buffercontaining 100 mmol L−1 KCl and 100 mmol L−1 AOT in isooctane. Proteins have back-extracted (82.7%) to a new aqueous phase containing 100 mmol L−1 pH 8.0 phosphatebuffers and 1,000 mmol L−1 KCl. No lipolytic activity was recovered after hydrophobicinteraction chromatography on a phenyl Sepharose column. The yield obtained for the overallprocess was 68% for enzyme activity, 26.4% for protein recovery with 810-fold purificationfactor, and a single protein band at 33,000 gmol−1 by SDS-PAGE [216].

A lipase from P. cyclopium MI was purified using ammonium sulfate precipitation, DEAE-Cellulose, DEAE-Sepharose, hydroxyapatite chromatographies, and gel filtration onCellulofine GC-700. The purification of the preparation was 1,380-fold and recovery yield27%. The molecular weight of the enzyme was estimated to be 35,000 gmol−1 fromSephadex G-100 chromatography [93]. Sztajer et al. [217] has purified a lipase enzymeproduced by P. simplicissimum by ammonium sulfate precipitation, phenyl-Sepharose CL-4B,Ultrogel AcA-54, and hydroxyapatite. The molecular weight of the enzyme was estimated at56,000 gmol−1 by SDS-PAGE with recovery of 20%. A lipase produced by a Penicilliumexpansum DSM 1994 was purified to 219-fold by cross-flow filtration, ammonium sulfateprecipitation, and phenyl-Sepharose with a specific activity of 558 Umg−1. The molecularweight of the homogeneous lipase has estimated as 25,000 gmol−1 by gel filtration and SDS-PAGE [218]. A lipase produced from Penicillium citrinum was purified by extraction andback-extraction using AOT reversed micelles in isooctane and phenyl-Superose column


[216]. The enzyme was purified 810-fold with a molecular mass of 33,000 gmol−1 by SDS-PAGE and recovery of 26.4%. Ferrer et al. [219] has purified a lipase of Penicilliumchrysogenum by ultrafiltration, phenyl-Sepharose, Mono Q HR 5/5, and PD-10 column onSephadex G-25. The purification of the preparation was 30-fold with a recovery yield of 44%and its molecular weight is 40,000 gmol−1. A lipase from P. cyclopium was purified to 590-fold by ammonium sulfate fractionation, Sephadex G-75, DEAE-Sephadex, and againSephadex G-75. The apparent molecular weight was 37,000 gmol−1 with a recovery yield of30%. Two types of lipases (lipases I and II) of Ustilago maydis ATCC 14826 was purified byLang et al. [220] using ion exchange chromatography on a Q-Sepharose column. Lipase I wasapplied to a butyl-Sepharose column with an overall yield of 12%. Lipase II was purified witha yield of 13.5% in DEAE-Sepharose and butyl-Sepharose columns. An extracellular lipasefrom Botrytis cinerea was purified to homogeneity using ammonium sulfate fractionation andsequential column chromatography on octyl-Sepharose CL-4B, Con A–Sepharose, andSephacryl S-200 columns. The purification of the preparation was 31-fold with a recoveryyield 21% and an apparent molecular mass of 60,000 gmol−1 on SDS-PAGE [221].

An extracellular lipase of G. candidum was purified by immune electrophoresistechnique [222]. Two lipolytic proteins (61,000 and 57,000 gmol−1) present in a SephadexG-100 fraction of extracellular lipase from G. candidum ATCC 66592 was separated usinghigh-performance liquid chromatography [223]. A lipase from G. candidum was purified byethanol precipitation and chromatography on Sephacryl-200 HR, high-resolution anionexchange (Mono Q) and Polybuffer exchanger 94. In this procedure, two forms of lipasesfrom G. candidum waS obtained. Lipase I and lipase II are purified at 35-fold with 62%recovery in activity and 94-fold with 18% recovery in activity, respectively [224]. A lipaseproduced by G. candidum was purified 13.2-fold using ion exchange chromatography onQ-Sepharose column followed by hydrophobic interaction on phenyl-Sepharose CL-4Bcolumn. In this procedure, pure lipase was obtained [222, 225]. One lipase was probablyremoved during hydrophobic interaction chromatography as 50% of lipase activity has beenlost in this step [226]. Shimada et al. [227] have purified a lipase produced by Fusariumheterosporum by SP-Sephadex chromatography, gel filtration, and isoelectric focusing withrecovery yield of 38%. The lipase has a monomeric protein with a molecular weight of31,000 gmol−1 estimated by SDS-PAGE.

The extracellular lipase from Fusarium sp. YM-30 was purified by a procedureinvolving ultrafiltration, ammonium sulfate precipitation, and DEAE-Toyopearl 650M,CM-Toyopearl 650 M, and Butyl-Toyopearl 650 M column chromatography. The purifiedlipase was homogeneous with 12,000 gmol−1 of molecular mass by SDS-PAGE, with highspecificity for mono and diacylglycerols, but low towards triacylglycerols [228]. Anextracellular lipase produced by Penicillium candidum has been purified 37-fold usingOctyl-Sepharose CL-4B and DEAE-Sephadex columns. The enzyme has been shown to bea monomeric protein of 29,000 gmol−1 molecular weight [229]. H. lanuginosa (now T.lanuginosus) produces an extracellular lipase which has been purified by means of acetoneprecipitation and successive chromatographic steps on Sephadex G-75, DEAE-SepharoseCL-6B, and hydroxyapatite columns [230]. The enzyme was purified about 150-fold andyield of 15% with single protein band on polyacrylamide gel electrophoresis. Its molecularweight was estimated as 39,000 gmol−1 on both SDS-PAGE and gel filtration on SephadexG-100, suggesting that the enzyme is a monomer. Taylor [231] has partially purified H.lanuginosa lipase by adsorption to acrylic microporous membranes in a pleated capsulefilter. The lipase has completely recovered by desorption at pH 9.0 (0.2 M Na2CO3),concentrated by ultrafiltration and freeze drying. About a quarter of the activity of theoriginal lipase remained in the freeze-dried powder. The preparation was further purified by


chromatography on Sephadex G-75. Boominathan et al. [232] has studied the effect ofmolecular weight of poly ethylene glycol (PEG) on purification of lipase of H. lanuginosaand reported that the ratio of desired protein to total protein has been improved by usinglower average molecular weight PEG with varying pH.

Gene Cloning

Genomic DNA encoding Lipase I is cloned from R. niveus IFO4759. For expression of thisgene in S. cerevisiae, DBY746 was transformed with YEp352PLipS, which had the clonedlipase gene under the control of a PGK promoter. This strain secreted the lipase at a highlevel (350 UmL−1). The strain ND-12B was produced by a mating of DBY746, harboringYEp352PLipS, and NA74-3A, and dissection of asci. This new strain secreted the lipase upto 530 UmL−1. Moreover, the lipase was produced most efficiently in a medium containingbacto-yeast extract, soy peptide, and sucrose [233]. Cloning of a lipase gene from R. oryzaeDSM 853 was studied [234]. Apparently, the different lipase forms of Rhizopus sp.described in the literature result from different proteolytic processing and created from thesame gene. Epitope mapping reported with monoclonal antibodies directed against humanpancreatic lipase and different mutant lipases propose that the beta 50 loop from C-terminaldomain may be involved in the interaction of human pancreatic lipase with a lipid/waterinterface [235].

C. rugosa has an unusual codon usage that interferes with the useful expression of genesderived from this fungus in conventional heterologous hosts. C. rugosa lipase occurs insome different isoforms encoded by the lip gene family. Of these lipases, the isoformsencoded by the gene lip1 is the most abundant. The lip1 gene (1647 bp) was completelysynthesized with an optimized nucleotide sequence to simplify genetic manipulation andallow heterologous expression in C. rugosa. The synthetic gene was functionallyoverexpressed in P. pastoris, allowing for the production of the specific isoformrecombinant lipase at a level of 150 UmL−1 in the culture medium. The physiochemicaland catalytic properties of the recombinant lipase was compared with those of acommercial, nonrecombinant, C. rugosa lipase preparation containing the lipase isoforms[236].

C. rugosa lipase isoenzymes share ca. 40% and 30% sequence homology with lipases ofG. candidum and Y. lipolytica, respectively. The domain of sequence conservation occurs atthe N-terminal half of the protein. For the resolution of isoforms via heterologousexpression, the lip 1 gene, encoding the major C. rugosa lipase form, was expressed in C.maltose, a related fungus with the same codon usage as C. rugosa. A recombinant lipasewas thus produced and secreted in an active form in the culture medium medium uponengineering the 5′- and 3′-ends of the gene [237].

R. niveus lipase has a unique structure consisting of two noncovalently boundpolypeptides (A and B chains). To improve this lipase by protein engineering, Kohno etal. [238] have developed a more efficient expression system for producing the lipase in S.cerevisiae. The expression system used the strain ND-12 B and the multicopy plasmidpJDB 219. A. oryzae produces at least three lipolytic enzymes, L1, L2, and L3 (mono- anddiglycerol, and triglycerol lipase, respectively). Of these, the L3 lipase gene was cloned.Nucleotide sequencing of DNA and cDNA revealed that the L3 gene had an open readingframe of 954 nucleotides, including three introns of 47, 83, and 62 bp. The deduced aminoacid sequences of the L3 gene implied a protein of 254 amino acid residues, with a singlesequence of 30 amino acids and, in spite of the difference in substrate specificity was


homologous to a sequence of fungal cutinases. Three residues presumed to form thecatalytic triad, Ser, Asp, and His, are preserved. The cloned cDNA of the L3 gene wasexpressed in Eschirichia coli to encode a functional triglycerol lipase [239].

Shimada et al. [240] investigated on G. candidum lipase by cloning and sequencing twoclosely related lipase genes lipI and lipII. Then, Bertolini et al. [241] investigated thepolymorphism of the lipase genes from G. candidum strains [242]. Candida deformansCBS 2071 produces an extracellular lipase that was cloned by complementation in S.cerevisiae with the genomic library of C. deformans. Three members of the lipase genefamily CdLIP1, CdLIP2, and CdLIP3 was cloned and characterized. Each deduced lipasesequence has a Glu-His-Ser-Leu-Gly- (Gly-/Ala)-Ala conserved motif, eight cysteineresidues and encodes an N-terminal signal sequence. The matrix-assisted laser desorption/ionization data suggests that it is LIPI that produces the lipase. These lipases have verysimilar to lipases from the Y. lipolytica [243].

Stehr et al. [244] have reported expression pattern of multigene family of Candidaalbicans using reverse transcription polymerase chain reaction in experimental infectionsand in samples of patients suffering from oral candidosis. The findings illustrate thatindividual lipase genes are differentially regulated in a mouse model system. This reportindicated that the lipase gene expression profile depended on stage of infection rather thanon organ localization. The temporal regulation of lipase gene expression was detected in anexperimental model of oral candidosis.

Yan et al. [245] reported the lipase gene from Galactomyces geotrichum Y05 was clonedfrom both genomic DNA and cDNA sources. Nucleotide sequencing revealed that the gglgene has an ORF of 1,692 bp without any introns, encoding a protein of 563 amino acidresidues, including a potential signal sequence of 19 amino acid residues. The amino acidsequence of this lipase showed 86% identity to lipase of Trichosporon fermentans WU-C12. The lipase gene was subcloned into pPIC9K vector and overexpressed in P. pastorisGS115. Active lipase has accumulated to the level of 100 UmL−1 in the shake-flask culture,10.4-fold higher than the activity of the original strain (9.6 UmL−1). The purified lipaseexhibited some properties of significant industrial importance, such as pH and temperaturestability, wide organic solvent tolerance, and broad hydrolysis on oils. Yu et al. [17] havestudied the cloning and expression of the lipase gene from R. chinensis in P. pastoris andcharacterization of the recombinant lipase. The lipase gene without its signal sequence havecloned downstream to the alpha-mating factor signal and expressed in P. pastoris GS115under the control of AOX1 promoter. The amino terminal analysis showed that the37,000 gmol−1 protein has the mature lipase (30,000 gmol−1) attached with 27 amino acidof the carboxy terminal part of the prosequence (r27RCL). The pH and temperatureoptimum of r27RCL and mRCL have pH 8.5 and 40 °C, and pH 8 and 35 °C, respectively.The chain length specificity of r27RCL and mRCL showed highest activity toward p-nitrophenyl hexanoate or glyceryl tricaproate (C6) and p-nitrophenyl acetate or glyceryltriacetate (C2), respectively. This property is quite rare among lipases and gives this newlipase great potential for use in the field of biocatalysis.

Lip2 lipase from Y. lipolytica is a very promising lipase with several potentialapplications such as resolution of racemic mixtures and synthesis of fine chemicals. Thispotential is impeded by a very low thermostability for temperatures higher than 40 °C.Saturation mutagenesis experiment at position 244 demonstrated that the presence of acysteine at this position was dependable for the thermal denaturation. It has demonstratedthat WT Lip2 and the thermostable variant are both inactivated through aggregationmechanisms. The WT lipase, rapid intermolecular disulphide bridge interchanges triggeredby the free cysteine 244 mediates aggregation. With the variant C244A, aggregation still


occurred but much slower than for the WT enzyme and mostly driven by hydrophobicforces [246].

Adachi et al. [247] have developed the A. oryzae whole-cell biocatalyst whichcoexpresses F. heterosporum lipase (FHL) and mono- and diglycerol lipase B in the samecell for biodiesel production. Screening using two selectable markers (sC and niaD)reported in the best transformant. FHL gene was integrated into A. oryzae chromosomeusing sC selection marker and mdlB was integrated using niaD selection marker.

Mathematical Modeling

Mathematical modeling in the fermentation process is an important tool for optimum designand operation of bioreactors to scale-up lipase production. The cost procured in predictingfermentation conditions at larger scale eases with modeling. Models in bioreactor can be oftwo types: kinetic models and transport models. A kinetic model describes how themicroorganisms are influenced by different process parameters, while the transport modeldescribes the mass and heat transfer within the bioreactor fermentation process.

A simple structured mathematical model coupled with a methodology to estimate statevariables and parameters has been reported for C. rugosa batch and continuous lipaseproduction. Process modeling has carried out using Advanced Continuous SimulationLanguage (ACSL). Model parameters was determined and validated with satisfactoryresults. For estimation of the best strategy to improve lipase productivity, differentsimulations of batch, fed-batch, and continuous cultures was proposed. The maximumenzyme productivity was predicted in a continuous culture at moderate dilution rates andsubstrate feeding of 8 gL−1. However, the highest predicted lipase activity was reached infed-batch cultures with a prefixed substrate feeding in order to maintain constant at optimalvalues of substrate/biomass or the specific growth rate. The fed-batch process mode wasmatched with the simulation results [248]. Freire et al. [249] developed a mathematicalmodel that describes time–course variations of extracellular lipase activities for the batchfermentation of P. restrictum strain isolated from soil and wastes of a Brazilian babassucoconut oil industry. The batch process was modeled by an unstructured model, consideringdependent variables like cells, fat acid, dissolved oxygen concentrations, lipase activities,and cell lysate concentration. The model was able to predict the effects of some operationvariables such as air flow rate and agitation speed. The mathematical model was validated.As biomass concentration has antagonistic effects on lipase activity, a typical optimizationscheme need to be developed in order to minimize these deleterious effects whilemaximizing lipase activity.

The Plackett–Burman statistical experimental design was developed to evaluate thefermentation medium components for the production of lipase by C. rugosa in submergedfermentation and an unstructured kinetic model to simulate the experimental data. Thelogistic model, the Luedeking–Piret model, and the modified Luedeking–Piret model hasbeen found to be suitable for efficient prediction of cell mass, lipase production, andglucose consumption with high correlation coefficients (R2). The estimated values of theLuedeking–Piret model parameters, α and β, found that the lipase production by C. rugosais growth-associated [99]. Boareto et al. [250] proposed an efficient hybrid neuralphenomenological model (HNM) for the lipase production process by C. rugosa, withthe bioreactor in batch and fed-batch operation. The experimental data are usedcorresponding to fed-batch operation with constant substrate feed rate. Artificial neuralnetworks were trained to represent the aqueous and intracellular lipase activity which was


further associated with a reduced version of the mechanistic model of the proposed HNM.When compared to the experimental data, the HNM exhibited higher accuracy. The HNMcan be used in process monitoring using online measurements of CO2 and substrate feedrate to infer enzyme activities and also substrate and biomass concentrations. A simplestructured mathematical model was developed [185] coupled with a methodology toestimate biomass amounts, specific growth rate, and substrate amounts and applied to theproduction of lipase by C. rugosa in batch, fed-batch, and continuous operations with atenfold increase in productivity relative to batch operation.

Conclusions

Critical analysis of the current literature shows that fungal lipases are one of the mostproduced enzymes. Its importance is increasing in several industries, such as food, dairy,detergent, pharmaceutical, etc. This review presents work of researchers for screening ofnew fungal lipase-producing microorganisms and on the optimization of the mediumcomposition and operational variables. The lipase production was developed bybioprocesses, mainly using submerged fermentation such as the screening of high lipaseproducers, successful substitution of synthetic media by agro-industrial residues, scale-upof process, different operation modes, and the use of mathematical models as a tool forprocess optimization. The screenings of fungal microorganism producers of lipases havesatisfactory results. The use of agro-industrial residues as substrates for lipase productionfavors the reduction of production cost. However, systematic investigation is required. Thefed batch mode has provided the good results in terms of lipase activity and productivity. Inall cases in this review, the feed rate is not optimized. An alternative optimization tool suchas the dynamic optimization can be used. The application of dynamic optimization toolmakes it possible to find the optimal profile of feeding rate that maximizes the productivity.The use of optimization tool requires a mathematical model to represent the process in asatisfactory and reliable way. SSF process might open new opportunities for the lipaseproduction since agro-industrial residues with low cost can be used as substrates with highproductivity as reported by numerous articles cited in this review. However, SSF is difficultto scale-up due to existing gradients in packed-bed reactor in temperature, pH, oxygen,substrate, inoculums, and moisture.

In addition, simple and efficient purification process is hydrophobic interaction chromatog-raphy. Lipase-based processing has promising future; however, the rate of development is slow.Factors posing limitations include a relatively high cost of lipase and a lack of enzymes withoptimal range of catalytic specificities and properties required in the different applications.

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Overview of Fungal Lipase: A Review

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