Table of Contents Author Guidelines Submit a Manuscript
BioMed Research International
Volume 2014 (2014), Article ID 108913, 11 pages
Research Article

Lipolytic Potential of Aspergillus japonicus LAB01: Production, Partial Purification, and Characterisation of an Extracellular Lipase

1Cell Signaling, Nanobiotechnology and Enzymology Laboratory, Federal University of Minas Gerais, 31270-901 Belo Horizonte, MG, Brazil
2Laboratory of Applied Microbiology, Federal University of Minas Gerais 31270-901, Belo Horizonte, MG, Brazil

Received 16 May 2014; Revised 31 August 2014; Accepted 2 September 2014; Published 29 October 2014

Academic Editor: Yunjun Yan

Copyright © 2014 Lívia Tereza Andrade Souza et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


Lipolytic potential of Aspergillus japonicus LAB01 was investigated by describing the catalytic properties and stability of a secreted extracellular lipase. Enzyme production was considered high under room temperature after 4 days using sunflower oil and a combination of casein with sodium nitrate. Lipase was partially purified by 3.9-fold, resulting in a 44.2% yield using ammonium sulphate precipitation (60%) quantified with Superose 12 HR gel filtration chromatography. The activity of the enzyme was maximised at pH 8.5, and the enzyme demonstrated stability under alkaline conditions. The optimum temperature was found to be 45°C, and the enzyme was stable for up to 100 minutes, with more than 80% of initial activity remaining after incubation at this temperature. Partially purified enzyme showed reasonable stability with triton X-100 and was activated in the presence of organic solvents (toluene, hexane, and methanol). Among the tested ions, only Cu2+, Ni2+, and Al3+ showed inhibitory effects. Substrate specificity of the lipase was higher for C14 among various p-nitrophenyl esters assayed. The and values of the purified enzyme for p-nitrophenyl palmitate were 0.13 mM and 12.58 umol/(L·min), respectively. These features render a novel biocatalyst for industrial applications.

1. Introduction

Aspergillus japonicus is amenable for biotechnological analysis due to its phytase [1], cellulose [2], pectinase [3], xylanase [4, 5], and beta-fructofuranosidase [6] production capacity. Furthermore, the production of fibrolytic enzymes is well described; however, their lipolytic potential remains unknown. Lipases (triacylglycerol hydrolases, EC. are a class of serine hydrolases that belong to the hydrolase super family. Depending on the water activity and the tolerance of organic solvents, lipases can act as modifying triacylglycerides by catalysing hydrolysis and different synthetic reactions, such as esterification, interesterification, and transesterification, with different specificities, efficiencies, and environmental compatibilities [7]. Interestingly, direct correlations between the enzyme activities in aqueous or organic media are lacking. Thus, some lipases have significant potential for application in oil and fat hydrolysis, but almost none have been applied in industrial processes, such as biodiesel production [8]. Reaction versatility is an important feature of this enzyme and is a subject to be investigated in the search for new lipases.

Lipases are regarded as the most important enzymes for biocatalysis in organic media and have been industrially used as lipid stain digesters in detergent formulation, the preparation of various products, and aromas in the food industry, structured lipids synthesis, leather processing, enantioresolution esters for chemical and drug intermediates, biodiesel production, and the treatment of waste products rich in oil [9].

Most industrial microbial lipases are derived from fungi, especially the Aspergillus genus [10]. Over the years, efforts have been directed to describe lipases from a variety of species, including A. nomius [11], A. niger [12], A. carneus [13], A. repens [14], A. terreus [15, 16], A. oryzae [17], and A. wentii [18]. In general, lipases obtained from Aspergillus are promising for their catalytic properties, such as thermal stability, specificity, stereospecificity, tolerance of organic solvents, and bulk production [10].

Lipases have emerged as leading biocatalysts, with less than 5% of participation in the global market; the market is expected to grow over the coming years, mainly due to the application of enzymes in the biofuel industry (biodiesel). Due to the increase in the biotechnological importance and potential of enzymes to contribute to the multibillion dollar unexploited bioindustry, a wide screening strategy for extra- and intracellular lipases that produce microorganisms has been evaluated. Previous studies conducted by our research group demonstrated that the isolated A. japonicus LAB01 was a potential lipase source based on both detection assays of lipase activity in solid and liquid media (data not shown). The present work aims to explore the A. japonicus LAB01 lipolytic activity by describing the production, purification, and biochemical characterisation of an extracellular lipase.

2. Materials and Methods

2.1. Microorganism and Extracellular Lipase Production by A. japonicus LAB01 in Shaker Flask Culture

The strain used in this work (A. japonicus LAB01) was isolated from urban matured solid compost waste obtained from Coimbra (Brazil, MG) and was kindly provided by the Laboratory of Biochemical Analysis/BIOAGRO/UFV (Brazil). This microorganism was maintained in glycerol and potato dextrose agar (PDA) slants under refrigeration (4°C). Cells were grown on basal medium containing the following (g/L): casein 1.0, NaNO3 1.0, K2HPO4 1.52, MgSO4 7H2O 0.52, and KCl 0.52, adjusted to pH 6.0, and supplemented with sunflower oil 1.0% (v/v). Cells were grown with shaking at 30°C (200 r.p.m.) for 96 h in 500 ml Erlenmeyer flasks containing 100 ml medium. The inoculum was prepared by transferring eight discs (0.6 cm in diameter) from PDA plate, after growth of the fungus for 96 h at 30°C. To test the effect of immobilised cells on the enzyme secretion, 1 gram of 6 mm cubes reticulated polyurethane foam was added to each flask before autoclaving and the fermentations were incubated as described above. These flasks were incubated at 30°C for 96 h on a reciprocal shaker (200 oscillations/min). When the cells were immobilised, 1 gram of 6 mm cubes reticulated polyurethane foam was added to each flask before autoclaving. The cultivated cells were separated from the culture broth by filtration using Whatman qualitative paper (number 1). The supernatant was considered crude enzyme and was used for the analytical assay. The immobilised cells were washed with tap water and dried at room temperature.

2.2. Assay of Lipase Activity in Aqueous and Nonaqueous Medium

The lipase activity was measured with a modified spectrophotometric method using p-nitrophenyl palmitate (p-NPP) as a substrate. For the hydrolytic assay, the substrate solution was prepared by mixing 1 mL of solution A (90 mg of p-NPP dissolved in 30 mL 2-propanol) and 9 mL of solution B (90 mM Tris-HCl buffer (pH 8.0); 2.0% Triton X-100; and 0.2% gum arabic). The enzyme-substrate mixture was incubated at 37°C for 5 minutes, and the change in the absorbance was measured at 410 nm in kinetic mode using a microplate reader Varioskan flash and in special cases Shimadzu UV-160A (tests were carried out in temperature higher than 45°C). The molar extinction coefficient of p-nitrophenol (p-NP) was estimated to be 1.27 × 103 M−1·cm−1. One enzyme unit (U) was defined as the lipase activity that liberated 1 nmol of p-NP per millilitre per minute under the standard assay conditions. For the transesterification assay, methanol and ethanol were used as acyl donors according to Teng and Xu [19].

2.3. Lipase Partial Purification

The fermented broth was filtered to obtain a clear supernatant, which was concentrated using ammonium sulphate precipitation at 60% saturation and 4°C with constant stirring overnight. The precipitated protein was collected by centrifugation at 10,000 ×g per 15 minutes and dissolved in a minimum volume of 20 mM Tris-HCl buffer at pH 8.0. The concentrated sample was applied to a Superose 12 HR 10/30 column coupled with the FPLC (fast protein liquid chromatography) system preequilibrated with 20 mM Tris-HCl buffer and pH 7.9 and a flow rate of 30 mL/h. The column was then washed with the same buffer until the flow-through fractions showed no lipase activity. The active fractions were pooled and concentrated by freeze-drying. This concentrated fraction was stored at 20°C until further use. The protein content was estimated using the Bradford reagent as previously described [20, 21].

2.4. Gel Electrophoresis and Zymography

The lipase molecular mass was determined by SDS-PAGE using a Laemmli system [22] with 12.5% acrylamide. The proteins were stained with a silver stain [23, 24]. For the zymographic analysis, the gel was incubated at 4°C with 100 mL of Tris-HCl buffer (pH 7.9, 20 mM) plus serum albumin (1% w/v) after running to promote protein folding. The gels were incubated for at least 4 hours with solution replacement every 30 minutes. The zymographic analysis was performed based on the lipase hydrolytic and synthetic activity using tributyrin [25], oleic acid, and dodecanol as substrates [26].

2.5. Partial Purified Lipase Biochemical Characterisation

All tests to characterise the lipases were conducted in triplicate using the enzyme pool obtained from the purification route described above. Methodologies were designed according to enzymatic characterisation studies employed for lipases from others sources [13, 2731].

2.5.1. Storage Stability of Crude Enzyme Determination

The storage stability of crude lipase was examined by conserving the cell-free culture supernatant at different temperatures: freezer (−20°C), ultrafreezer (−80°C), fridge (4°C), and room temperatures (22–26°C). The lipase activity was measured every 7 days with a standard pNPP-hydrolysing assay.

2.5.2. pH Effect on Enzyme Activity and Stability

The pH stability was studied by incubating the purified enzyme for 90 min at 30°C in various buffers: 100 mM citrate phosphate buffer (pH 3.0–6.0), 100 mM sodium phosphate buffer (pH 7.0), 100 mM Tris-HCl buffer (pH 8.0-9.0), and 100 mM glycine-NaOH buffer (pH 10.0). The residual hydrolytic activity was then assessed under standard assay conditions using p-NPP as substrate at 37°C and pH 8.0. The optimal pH for the enzyme was determined by measuring the lipase activity using different pH levels for solution B (pH 7–9.5) at 37°C.

2.5.3. Temperature Effect on Enzyme Activity and Stability

The optimal enzyme temperature was determined by assessing the enzymatic kinetics described above at temperatures ranging from 20 to 60°C. The results of this assessment were used to investigate the enzyme thermal stability in that condition by incubating the purified enzyme for 12 h followed by measuring the residual activity.

2.5.4. Organic Solvents, Ions, and Detergents Effect on Lipase Stability/Activity

Various effectors, including pure or water-mixed organic solvents (methanol, ethanol, isopropanol, acetone, acetonitrile, toluene, and hexane), ionic solutions prepared in 200 mM Tris-HCl (pH 8.0) buffer (K+, Li+, Mg2+, Ca2+, , Ni2+, Na+, Cu2+, and Al3+), and detergents (1% w/v) (SDS, Tween 80, Triton X-114, and Triton X-100) were added at a proportion of 1 : 1 (v/v) to the purified lipase and incubated in an Eppendorf ThermoMixer (150 rpm) for 90 minutes at 30°C. The lipase activities of the sample without any organic solvent, ions, or detergent were taken as the control (100% of activity). The residual activity was measured with a p-NPP spectrophotometric assay at 37°C and pH 8.0.

2.5.5. Substrate Specificity

The enzyme substrate specificity was investigated by using p-nitrophenyl fatty acid esters of different chain lengths (p-nitrophenyl acetate (C2), p-nitrophenyl butyrate (C4), p-nitrophenyl octane (C8), p-nitrophenyl laurate (C10), p-nitrophenyl myristate (C14), and p-nitrophenyl palmitate (C16) as substrates). The lipase activity was assayed under standard conditions and various preparations of solution A.

2.5.6. Kinetics Parameters Determination

The apparent kinetic parameters (KM, , and ) of the partial purified lipase were calculated using a Michaelis-Menten hyperbola direct regression obtained experimentally with the SigmaPlot (v.10) software. The assays were carried out according to the standard reaction using p-nitrophenyl palmitate (C16) as substrate at concentrations varying from 0 to 1.4 mM.

2.6. Vegetable Oils Hydrolysis

The hydrolytic activity of crude extracellular lipase from A. japonicus LAB01 was determined on emulsified vegetable oils (corn, sunflower, soybean, olive, canola, pequi, almond, macauba, and sesame), according to Soares et al. [32]. The formed fatty acids were titrated with 20 mmol·L−1 sodium hydroxide solution in the presence of phenolphthalein as indicator. One international unit (IU) of activity was defined as the amount of enzyme that liberates 1 μmol free fatty acid per minute under the assay conditions.

3. Results and Discussion

3.1. Comparison of Extracellular Lipase Production between Suspension and Immobilised Cells

Supernatants collected after culturing cells in suspension and immobilised into polyurethane foam significantly differed in extracellular lipase activity. For the cell suspension, the lipase activity on supernatant was more than three times greater than the activity obtained for immobilised cells, 28.04 ± 3.11 and 8.09 ± 0.82 (U/mL), respectively. The reduction in lipase secretion during fermentation using immobilised biomass has been attributed to changes in the cell morphology, which can strongly inhibit the secretion of lipase to the medium [33]. Hama et al. [33] showed that the extracellular lipase activity of Rhizopus oryzae immobilised cells was approximately half that of suspension cells. Conversely, the intracellular lipase activity of immobilised cells was much higher than that of the suspension cells. Cell immobilisation occurred as a consequence of natural fungi growth, and the liquid media did not contain mycelia mass that had exited the support. The fungi grew into the pores, forming a web biomass that strongly adhered to the support and showed intracellular lipase activity (3.69 ± 0.5 U/g). This result is interesting because it reveals that the lipolytic potential of our strain is not limited to enzyme secretion. The use of intracellular lipases as immobilised biomass has become a promising alternative to catalyse organic reactions, especially for biodiesel production.

3.2. Extracellular Lipase from A. japonicus LAB01 Acting in Organic Medium Performance

To test the catalytic efficiency of extracellular lipase from A. japonicus LAB01 to act in an organic medium, the supernatant was concentrated and completely dried, which prevented a parallel hydrolysis reaction. The enzyme could catalyse the pNPP transesterification using anhydrous methanol and ethanol as the acyl donor group, with 23.60 ± 0.93 and 30.336 ± 2.60 (U/mL) of enzymatic activity, respectively. The enzyme more efficiently catalysed the hydrolysis reaction with an activity of 199.5 ± 10 (U/mL). In general, the catalytic activity displayed by enzymes in neat organic solvents is far lower than in water [19, 20, 30, 34]. This difference has been attributed to the enzyme’s decreased flexibility, denaturation by contact with the organic solvent, and diffusion effects [30]. This difference could also be due to the drying step effect. For example, lyophilisation can significantly change the enzyme structure to reduce the enzymatic activity in organic solvents [34]. The obtained data agree with those observed for lipases from Pseudomonas aurantiogriseum. This enzyme exhibited an activity of 312.3 (U/mg) and 13.6 (U/mg) on the pNPP substrate in organic and aqueous media, respectively [30]. Commercial lipases from Candida rugosa, Penicillium sp., and pig pancreas demonstrated the absence of p-NPP transesterification in the presence of ethanol [19]. However, the enzymes effectively catalysed the substrate hydrolysis. In fact, the hydrolytic and synthetic activities of lipases do not directly correlate. These preliminary results demonstrated the versatility of reactions with lipase from A. japonicus LAB01 and suggest that this enzyme could be applied in the transesterification catalysis of natural substrate (oil and fat) to produce biodiesel.

3.3. Lipase Partial Purification

A. japonicus LAB01 lipases were purified using (NH4)2SO4 selective precipitation (60%) followed by gel filtration chromatography (Table 1). After two steps of purification, the lipase exhibited 3.5 × 104 U·mg−1 specific activity and 2.91-fold purification with an overall yield of 44.2%. The protein purification step profile showed a strong protein band with a molecular mass between 25 and 35 kDa estimated by SDS-PAGE (Figure 1). The zymographic analysis using tributyrin, oleic acid, and dodecanol as substrates revealed hydrolytic and synthetic activities based on bands that coincided with the migration of the principle protein (Figure 2). The partially purified lipase molecular mass was estimated to be approximately 25 kDa on SDS-PAGE. Most of the known Aspergillus sp. lipases reportedly have molecular masses in the range of 29−70 kDa [7, 10]. Another lipase from A. japonicus was purified by combining anion exchange (Q-Sepharose) and gel filtration (Sephadex G-100) chromatography by 3.44-fold with a 4.53% yield and a specific activity of 480 U/mg protein. The molecular weight of the enzyme was 43kDa according to SDS-PAGE [35]. Other lipase isoforms also differ in their molecular mass, such as those observed on Aspergillus niger. Two lipases from A. niger MZKI A116 were isolated by acetone precipitation followed by ion-exchange chromatography. The molecular weights were 43 kDa and 65 kDa, with isoelectric points of pH 4.1 and 4.2, respectively [36]. Recently, a novel lipase produced by the same fungus was found [37]. An extracellular lipase from the Aspergillus niger NCIM 1207 strain was purified by homogeneity using ammonium sulphate precipitation followed by phenyl-sepharose and Sephacryl-100 gel chromatography to yield a monomeric protein of 32.2 kDa.

Table 1: Summary of purification of extracellular lipase from A.  japonicus LAB01.
Figure 1: SDS-PAGE of purified lipase from A. japonicus LAB01. Lane 1: molecular mass markers; Lane 2: purified lipase after (NH4)2SO4 precipitation; Lane 3: purified lipase after gel filtration.
Figure 2: Zymographic analysis after SDS-PAGE gel electrophoresis of lipase from A. japonicus LAB01. Lane 1: lipase activity towards tributyrin; Lane 2: lipase activity towards oleic acid and dodecanol; Lane 3: molecular mass markers.

3.4. Biochemical Characterisation of Lipases from A. japonicus LAB01
3.4.1. Crude Lipase Storage Stability Determination

To verify the storage stability of the crude extract that contained extracellular lipase from A. japonicus LAB01, the supernatant was stored under the following conditions: refrigerator (4°C), room temperature, freezer (−20°C), and ultrafreezer (−80°C), and the enzyme activity was measured weekly for 1 month. As shown in Figure 3, the enzyme retained at least 40% of its initial activity under the tested temperature conditions. The best supernatant storage condition was found to be in the ultrafreezer (−80°C). The enzyme retains approximately 80% of its initial activity (%) after 4 weeks of storage. Good storage stability for lipase from A. japonicus LAB01 is a prerequisite for laboratory bench scale tests and for its future industrial application. Similar results were found for an extracellular Lactobacillus plantarum lipase, which was stable at −80°C for 4 weeks, retaining 90% of its initial activity [38].

Figure 3: Storage stability of crude lipases from A. japonicus LAB01. Freezer (−20°C) (diamond), ultrafreezer (−80°C) (triangle), fridge (4°C) (circle), and room temperature (square).
3.4.2. Effect of pH on Lipase Stability and Activity

The lipase stability was evaluated in the pH range of 3.0 to 10.0 using different buffers at the same concentration (100 mM) (Figure 4). The enzyme remained stable from pH 5.0 to 9.0 during preincubation for 90 min at 30°C, where the enzyme retained at least 80% of its initial activity. Furthermore, increasing or decreasing the pH values dramatically reduced the enzyme stability. The A. japonicus LAB01 lipase was more active at slightly alkaline pH values, and the optimal pH was 8.5, which is similar to the values reported for lipases from A. carneus [13]. The purified enzyme from our strain did not show differences in the optimum pH and pH stability; consequently, the lipase could be considered a resistant alkaline lipase. Alkaline lipases are required for applications in the detergent industries, suggesting a possible use in this area. The majority of the lipases of Aspergillus origin showed maximum activities in neutral or near neutral pH values (6.0–7.5), such as those obtained from A. japonicus [35], A. niger MTCC 2594 [39], A. nidulans [40], A. niger F044 [41], A. awamori [42], A. niger MYA135 [27], and A. terreus [43]. A. niger NCIM 1207 lipase, which exhibited an optimum acidic pH of 2.5, was an exception to this rule [37].

Figure 4: Effects of pH on the stability (a) and activity (b) of A. japonicus LAB01 lipase. (a) Partially purified lipase samples were incubated for 90 min at 30°C in the presence of different buffers (100 mM): citrate phosphate (pH 3.0–6.0), sodium phosphate (pH 7.0), Tris-HCl (pH 8.0-9.0), and glycine-NaOH (pH 10.0), and the residual activity was measured using pNPP at pH 8.0 and 37°C. (b) Partially purified lipase samples were assayed in buffers B from pH 7.0 to 9.5 using pNPP as substrate at 37°C.
3.4.3. Temperature Effect on Lipase Activity and Stability

The purified lipase was active in the temperature range of 20–45°C, with maximal activity at 45°C. Its activity sharply decreased above the optimum temperature (Figure 5). The characteristics of A. japonicus LAB01 lipase were not consistent with those of others found in Aspergillus genus, including A. niger MTCC 2594 [39], A. carneus [13], and A. niger MYA135 [44]. In all of these cases, the optimum temperature was 37°C, and the enzymes were not very active at temperatures above 40°C. The recombinant A. fumigatus lipase, whose optimal temperature was 65°C, constitutes an exception. However, this enzyme was not stable at this temperature, losing almost 100% of its activity after 1 h of incubation [29]. The thermal stability profile of A. japonicus LAB01 lipase indicated that this enzyme was stable at 45°C for at least 3 h, retaining 70% of its original activity. The high thermal stability of the enzyme is similar to that found for the thermostable lipases obtained from R. homothallicus, which had a half-life equal to 0.72 hours at 50°C [45]. The thermal stability of lipase from our strain indicates its potential application in the modification of vegetable oils, mainly oils that have high melting points, which are solid at room temperature. In general, fungal lipases are not stable at temperatures above 40°C. Some studies have reported lipases with moderate thermostability, as observed for P. wortmanii lipase, which retained 90 and 55% of its initial enzymatic activity after 60 min of incubation at 45 and 50°C, respectively [46]. P. cyclopium lipase was stable for 1 hour when incubated at 30°C and only lost 10% of its activity after 1 hour of incubation at 35°C [47]. P. crustosum lipase lost 75% of its enzymatic activity after incubation for 1 hour at 45°C [48].

Figure 5: Effects of temperature on stability (a) and activity (b) of the partially purified A. japonicus LAB01 lipase. (a) Temperature stability profile was determined by measuring the residual activity at pH 8.0 and 37°C after incubating the lipase at 45°C for 720 min. (b) Optimal temperature was determined by measuring the enzyme activity at different temperatures at pH 8.0.
3.4.4. Substrate Specificity

The specificity of lipases is related to the carboxylic acid ester type and sets them apart from conventional chemical catalysts, especially for their application in structured triacylglycerol synthesis, chemical reaction intermediates, and resolution of chiral compounds. Therefore, this property is of great industrial importance and depends on the enzyme source. The specificity of the partially purified A. japonicus LAB01 lipase for chain length was evaluated based on the initial hydrolysis rate, employing acyl esters of p-nitrophenyl (pNP) with carbon chain sizes ranging from C2 to C16: acetate (C2, pNPA), butyrate (C4, pNPB), caprylate (C8, pNPC), decanoate (C10, pNPD), laurate (C12, pNPL), myristate (C14, pNPM), and palmitate (C16, pNPP). The specific profile showed that the enzyme activity positively correlated with the length of the substrate carbon chain (Table 2). The enzyme showed little activity on short-chained acyl esters, such as pNPA (C2) and pNPB (C4), and had the highest activity on pNPM (C14) (Table 2). The preference for pNPM (C14) substrate was also observed for lipase from Serratia marcescens ECU1010 [49]. Conversely, another lipase purified from A. japonicus exhibited a preference for the pNPP (C16) substrate, and less than 80% residual activity was found on pNPM [35]. Most of the described microbial lipases are more active in the cleavage of long or medium chain fatty acid substrates, as noted for lipases mentioned before as well as lipases from A. carneus, Staphylococcus aureus, and Penicillium aurantiogriseum, which showed a preference for pNPD substrate (C10) [13, 30, 50]; A. niger MYA lipase prefers pNPO (p-nitro substrate) (C18) [27]; P. crustosum lipase prefers pNPL (C12) [48]. However, a new type of lipase gene, which was cloned from the genomic DNA of A. fumigatus (CGMCC 2873), showed extremely high hydrolytic activity toward p-short-chain nitrophenyl esters, such as pNPA (C2) [29].

Table 2: Substrate specificity of A. japonicus LAB01 lipase for various acyl esters of p-nitrophenyl (pNP).

3.4.5. Ions Effect on Lipase Stability

Lipases are known not to require cofactors to act; however, metal ions significantly affect the stability of lipases. Some metal ions interact with the enzyme surface charge, affecting the ionization of some amino acid residues, changing the enzyme conformation, and rendering it less stable and/or altering its activity. This process is known as ion toxicity [51]. The behaviour of A. japonicus LAB01 lipase in the presence of some ions was investigated using chloride or sulphate salts of each ion at a concentration of 10 mM in Tris-HCl buffer (pH 8, 200 mM) at 30°C for 90 minutes. The results showed that K+, Na+, Ca2+, Mg2+, , and Li+ ions slightly increase the lipase activity up to 12% or exerted no effect on the activity, whereas lipase activity was inhibited by Cu2+, Ni2+, and Al3+ (Table 3). Enzymes, such as lipases, need to be stable at high salt concentrations in some cases, such as when a lipase is used in a waste processes, like the continuous removals thin layers of fats and oils from the surface of aerated tanks to permit oxygen transportation. Wastewater sources and their ion contents constitute another variable in this process. Furthermore, ion-exchange chromatography is a common step in purification methods used in lipase purification strategies, and adding specific ions to the buffer can increase the yield. Metal ions also affect lipase substrates; Hasan et al. [9] reported that these metals would form complexes with ionised fatty acids, changing their solubility and behaviour at the interfaces. The stability of the lipase depends on its source and even the isoforms. The addition of 2 mM metal ions in a 50 mM Tris-HCl buffer (pH 8.0) at 30°C for 0.5 h did not increase the activity of lipase from A. fumigatus. The enzyme was stable (residual activity > 95%) in Ca2+ and Ni2+. Conversely, Cu2+ and Zn2+ inhibited the lipase (residual activity < 50%) [29]. Incubation with 10 mM Ba2+ at 30°C for 30 minutes stimulated the activity of A. terreus var. africanus lipase by 52%, while Cu2+, Fe2+, Fe3+, Hg2+, and K+ completely inhibited the enzyme [43].

Table 3: Effect of ions on the activity/stability of the lipase from A.  japonicus LAB01.

3.4.6. Detergents Effect on Lipase Stability

Detergents have been used in some lipase activity assay protocols for immobilisation and biotransformation and may have both a positive and a negative effect on the enzyme activity, which depends on the concentration or detergent nature. As shown in Table 4, ionic surfactants, such as Tween 80 and Triton X-100 at 1% (v/v) decreased the lipase activity by 70 and 12%, respectively. Similar to lipases from A. fumigatus, which were unstable in SDS (0.1% v/v) [29], the A. japonicus LAB01 lipase was sensitive to the anionic surfactant SDS, which completely inhibited the lipase hydrolytic activity. The negative effects of detergents on the stability of A. japonicus LAB01 lipase can be attributed to the competition between the substrate and detergent for the adsorption site in the enzyme or distortion in the enzyme structure [52, 53]. The concentration of the detergent evaluated may also exert negative effects. Studying the stability of free and immobilised lipases from Pseudomonas fluorescens [53] showed that the activity/detergent concentration curve reached a maximum activity value and began to decrease as the detergent concentration increased, in many instances even below the initial value.

Table 4: Effect of detergents on the activity/stability of the lipase from A. japonicus LAB01.

Conversely, the lipase from Spirulina platensis was active in the presence of 0.1% Triton X-100 or 0.5 mM SDS, showing residual activities (%) of 121 and 103, respectively [54]. The large increase in the lipase activity, which was mainly observed for the immobilised form, can be attributed to the potential of these surfactants to break the lipase-lipase dimmers and alter the closed-open equilibrium of the individual lipase molecules [53, 55]. Moreover, the presence of detergents also improved the enantioselectivity of lipases. For example, the enantioselectivity of Pseudomonas fluorescens immobilised in glyoxyl agarose increased from 40 to more than 100 in the (±)-2-hydroxy-4-phenylbutyric acid ethyl ester hydrolysis when using 0.1% (v/v) hexadecyltrimethylammonium bromide (CTAB) [53].

3.4.7. Solvent Effect on Lipase Stability

Reaction media that consist of organic solvents can greatly expand the lipase-catalysed transformation repertoire, where numerous reactions might be used by synthetic and polymer chemists with high selectivity [34]. Furthermore, solvents affect the catalytic proprieties, such as the selectivity lipases, and increase the substrate solubility, eliminating mass-transfer barriers for substrates [56]. Hence, the stability and activity of lipases in solvents are considered an important investigation point. To study effects of organic solvent on A. japonicus LAB01 lipase, the enzyme was incubated with different solvents at 30°C for 90 min, and the residual enzymatic activity was determined at assay conditions. As shown in Table 5, the enzyme was activated in pure nonpolar solvents, showing 100% and 137% residual activity in toluene and hexane, respectively. Toluene has a strong antimicrobial potential; thus, it constitutes an alternative method to preserve A. japonicus LAB01 lipases during storage. The enhancement of enzyme activity due to solvents could be attributed to lipase disaggregation, some structural changes in the enzymes (maintaining the lipase on its open conformation), and even modification at the oil-water interface to facilitate enzymatic action [57].

Table 5: Effect of organic solvents on the activity/stability of the lipase from A. japonicus LAB01.

Conversely, the lipase molecule was unstable in pure polar solvents, completely losing its activity in acetone and acetonitrile. Regarding the solvents employed in biodiesel production, the enzyme was more stable in a methanol solution, mainly at 50% (v/v) concentration, than in ethanol and propanol. The effect of the solvent polarity does not explain this finding; rather, this effect is likely due to the similar sizes of the water and methanol molecules, which can decrease the distortion in the three-dimensional structure of the protein. The enzyme was sensitive to the increased alcohol concentration, almost losing activity at increasing alcohol concentrations up to 75% (v/v).

In general, hydrophobic solvents are usually superior to hydrophilic ones as enzymatic reaction media because the latter are likely to tightly strip the bound water (which is essential to the catalytic properties and structure) from the enzyme molecules [34]. In fact, lipases are diverse in their sensitivity to organic solvents, but polar solvents are generally agreed to be more destabilising than nonpolar solvents, as has been demonstrated for lipases from Staphylococcus sp. [58], Penicillium sp. DS-39 (DSM 23773) [59], and Idiomarina sp. W33 [60].

Yang et al. [56] studied the effect of organic solvents on the conformation of recombinant Candida antarctica lipase (CALA). Treating CALA with acetonitrile increased the α-helical content and decreased the random coil content. The changes in the secondary structure of the enzyme were mainly responsible for the variance of the catalytic activity, which allowed this enzyme to be used for organic synthesis and chiral resolution. However, no obvious relationships could be established for any distinctive change in activity caused by the types of solvents.

3.4.8. Kinetics Parameters Determination

The apparent kinetic parameters for partially purified lipase were obtained from a rectangular hyperbola equation using the SigmaPlot software (version 10.0) (Figure 6). It yielded apparent KM and values of 0.13 mM and 12.58 μM·min−1, respectively, for p-NPP as a substrate. Similar kinetic parameters were also found for A. awamori (0.13 mM) [61] and A. nidulans (0.28 mM) lipases [62].

Figure 6: Michaelis-Menten hyperbola plot for KM and values of A. japonicus LAB01 lipase in the presence of various concentrations of p-NPP.
3.5. The Application of Lipases to Catalyse Vegetable Oil Hydrolysis

Lipases from A. japonicus LAB01 could hydrolyse all commercial oils tested and activities were expressed as percent of activity against canola oil. The enzyme preferentially catalysed canola and almond oils, showing yields of 100 ± 0.84 and 83 ± 0.73%, respectively. For other oils evaluated, the yields (%) were: corn oil (62 ± 0.01), pequi oil (55 ± 0.42), sesame oil (52 ± 0.02), sunflower oil (52 ± 0.01), soybean oil (48 ± 0.42), and olive oil (38 ± 0.42). As mentioned above, the specificity of lipases depends on their source. Rice bran oil and sardine oil were effectively hydrolysed by Cryptococcus sp. S-2 lipases with 80.2 and 76% activity on triolein [63]. Rhizopus oryzae lipases more efficiently hydrolysed olive oil, with a yield of 48% against trioctanoin [31]. From a biotechnological point of view, the use of lipases in the oleochemical industry is adequate because is an energy-saving process and unsaturated fatty acids can be produced without oxidation [64].

4. Conclusion

The lipolytic potential of A. japonicus LAB01 was demonstrated based on its ability to produce extracellular and intracellular lipases. The extracellular lipase from A. japonicus LAB01 showed versatile and interesting catalytic properties: it may act in aqueous and nonaqueous media, exhibiting maximum activities at alkaline pH values and increased temperatures (45°C), and it is compatible with processes used to modify fats and oils. Furthermore, the enzyme showed good stability when placed in contact with organic solvents, salt, and detergents. Future developments in the optimisation of extra- and intracellular lipase production and biotechnological applications for this lipase will be interesting to follow.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.


The authors are grateful for CNPq (Conselho Nacional de Desenvolvimento Científico e Tecnológico), INCT de Nanomateriais de Carbono, FAPEMIG (Fundação de Amparo à Pesquisa do Estado de Minas Gerais), Rede Mineira de Toxinas com Ação Terapêutica, and CAPES for financial supports.


  1. A. Maller, A. R. L. Damasio, T. M. Silva, J. A. Jorge, H. F. Terenzi, and M. L. T. M. Polizeli, “Potential application in animal feed of phytase produced from agro-industrial residues by Aspergillus japonicus,” Journal of Biotechnology, vol. 150, p. 514, 2010. View at Google Scholar
  2. F. D. A. Facchini, A. C. Vici, V. R. A. Reis et al., “Production of fibrolytic enzymes by Aspergillus japonicus CO3 using agro-industrial residues with potential application as additives in animal feed,” Bioprocess and Biosystems Engineering, vol. 34, no. 3, pp. 347–355, 2011. View at Publisher · View at Google Scholar · View at Scopus
  3. M. V. Semenova, S. G. Grishutin, A. V. Gusakov, O. N. Okunev, and A. P. Sinitsyn, “Isolation and properties of pectinases from the fungus Aspergillus japonicus,” Biochemistry, vol. 68, no. 5, pp. 559–569, 2003. View at Publisher · View at Google Scholar · View at Scopus
  4. F. D. A. Facchini, A. C. Vici, V. M. Benassi et al., “Optimization of fibrolytic enzyme production by Aspergillus japonicus C03 with potential application in ruminant feed and their effects on tropical forages hydrolysis,” Bioprocess and Biosystems Engineering, vol. 34, no. 8, pp. 1027–1038, 2011. View at Publisher · View at Google Scholar · View at Scopus
  5. M. Wakiyama, K. Yoshihara, S. Hayashi, and K. Ohta, “An extracellular endo-1,4-β-xylanase from Aspergillus japonicus: purification, properties, and characterization of the encoding gene,” Journal of Bioscience and Bioengineering, vol. 109, no. 3, pp. 227–229, 2010. View at Publisher · View at Google Scholar · View at Scopus
  6. S. I. Mussatto, C. N. Aguilar, L. R. Rodrigues, and J. A. Teixeira, “Colonization of Aspergillus japonicus on synthetic materials and application to the production of fructooligosaccharides,” Carbohydrate Research, vol. 344, no. 6, pp. 795–800, 2009. View at Publisher · View at Google Scholar · View at Scopus
  7. R. Sharma, Y. Chisti, and U. C. Banerjee, “Production, purification, characterization, and applications of lipases,” Biotechnology Advances, vol. 19, no. 8, pp. 627–662, 2001. View at Publisher · View at Google Scholar · View at Scopus
  8. K. Tongboriboon, B. Cheirsilp, and A. H-Kittikun, “Mixed lipases for efficient enzymatic synthesis of biodiesel from used palm oil and ethanol in a solvent-free system,” Journal of Molecular Catalysis B: Enzymatic, vol. 67, no. 1-2, pp. 52–59, 2010. View at Publisher · View at Google Scholar · View at Scopus
  9. F. Hasan, A. A. Shah, and A. Hameed, “Industrial applications of microbial lipases,” Enzyme and Microbial Technology, vol. 39, no. 2, pp. 235–251, 2006. View at Publisher · View at Google Scholar · View at Scopus
  10. F. J. Contesini, D. B. Lopes, G. A. MacEdo, M. D. G. Nascimento, and P. D. O. Carvalho, “Aspergillus sp. lipase: potential biocatalyst for industrial use,” Journal of Molecular Catalysis B: Enzymatic, vol. 67, no. 3-4, pp. 163–171, 2010. View at Publisher · View at Google Scholar · View at Scopus
  11. M. M. R. Talukder, H. Z. S. Lee, R. F. Low, L. C. Pei-Lyn, D. Warzecha, and J. Wu, “Potential use of whole cell lipase from a newly isolated Aspergillus nomius for methanolysis of palm oil to biodiesel,” Journal of Molecular Catalysis B: Enzymatic, vol. 89, pp. 108–113, 2013. View at Publisher · View at Google Scholar · View at Scopus
  12. N. Z. Adham and E. M. Ahmed, “Extracellular lipase of Aspergillus niger NRRL3; production, partial purification and properties,” Indian Journal of Microbiology, vol. 49, no. 1, pp. 77–83, 2009. View at Publisher · View at Google Scholar · View at Scopus
  13. R. K. Saxena, W. S. Davidson, A. Sheoran, and B. Giri, “Purification and characterization of an alkaline thermostable lipase from Aspergillus carneus,” Process Biochemistry, vol. 39, no. 2, pp. 239–247, 2003. View at Publisher · View at Google Scholar · View at Scopus
  14. Y. Kaminishi, H. Tanie, and M. Kunimoto, “Purification and characterization of lipases from Aspergillus repens and Eurotium herbariorum NU-2 used in “Katsuobushi” molding,” Fisheries Science, vol. 65, no. 2, pp. 274–278, 1999. View at Google Scholar · View at Scopus
  15. R. P. Yadav, R. K. Saxena, R. Gupta, and W. S. Davidson, “Purification and characterization of a regiospecific lipase from Aspergillus terreus,” Biotechnology and Applied Biochemistry, vol. 28, no. 3, pp. 243–249, 1998. View at Google Scholar · View at Scopus
  16. R. Gulati, R. K. Saxena, and R. Gupta, “Fermentation waste of Aspergillus terreus: a promising copper bio-indicator,” Current Science, vol. 77, no. 10, pp. 1359–1360, 1999. View at Google Scholar
  17. K. Ohnishi, Y. Yoshida, J. Toita, and J. Sekiguchi, “Purification and characterization of a novel lipolytic enzyme from Aspergillus oryzae,” Journal of Fermentation and Bioengineering, vol. 78, no. 6, pp. 413–419, 1994. View at Publisher · View at Google Scholar · View at Scopus
  18. H. Chander, V. K. Batish, S. S. Sannabhadti, and R. A. Srinivasan, “Factors affecting lipase production in Aspergillus wentii,” Journal of Food Science, vol. 45, no. 3, pp. 598–600, 1980. View at Google Scholar
  19. Y. Teng and Y. Xu, “A modified para-nitrophenyl palmitate assay for lipase synthetic activity determination in organic solvent,” Analytical Biochemistry, vol. 363, no. 2, pp. 297–299, 2007. View at Publisher · View at Google Scholar · View at Scopus
  20. E. Lorençon, A. S. Ferlauto, S. De Oliveira et al., “Direct production of carbon nanotubes/metal nanoparticles hybrids from a redox reaction between metal ions and reduced carbon nanotubes,” ACS Applied Materials and Interfaces, vol. 1, no. 10, pp. 2104–2106, 2009. View at Publisher · View at Google Scholar · View at Scopus
  21. S. L. Da Silva, J. R. Almeida, L. M. Resende et al., “Isolation and characterization of a natriuretic peptide from crotalus oreganus abyssus (grand canyon rattlesnake) and its effects on systemic blood pressure and nitrite levels,” International Journal of Peptide Research and Therapeutics, vol. 17, no. 3, pp. 165–173, 2011. View at Publisher · View at Google Scholar · View at Scopus
  22. U. K. Laemmli, “Cleavage of structural proteins during the assembly of the head of bacteriophage T4,” Nature, vol. 227, no. 5259, pp. 680–685, 1970. View at Publisher · View at Google Scholar · View at Scopus
  23. V. Paschon, G. S. V. Higa, R. R. Resende, L. R. G. Britto, and A. H. Kihara, “Blocking of connexin-mediated communication promotes neuroprotection during acute degeneration induced by mechanical trauma,” PLoS ONE, vol. 7, no. 9, Article ID e45449, 2012. View at Publisher · View at Google Scholar · View at Scopus
  24. E. S. de Alvarenga, S. A. Silva, L. C. A. Barosa et al., “Synthesis and evaluation of sesquiterpene lactone inhibitors of phospholipase A2 from Bothrops jararacussu,” Toxicon, vol. 57, no. 1, pp. 100–108, 2011. View at Publisher · View at Google Scholar · View at Scopus
  25. R. Singh, N. Gupta, V. K. Goswami, and R. Gupta, “A simple activity staining protocol for lipases and esterases,” Applied Microbiology and Biotechnology, vol. 70, no. 6, pp. 679–682, 2006. View at Publisher · View at Google Scholar · View at Scopus
  26. M.-A. Kwon, H. S. Kim, D.-H. Hahm, and J. K. Song, “Synthesis activity-based zymography for detection of lipases and esterases,” Biotechnology Letters, vol. 33, no. 4, pp. 741–746, 2011. View at Publisher · View at Google Scholar · View at Scopus
  27. C. M. Romero, L. M. Pera, C. Olivaro, A. Vazquez, and M. D. Baigori, “Tailoring chain length selectivity of a solvent-tolerant lipase activity from Aspergillus niger MYA 135 by submerged fermentation,” Fuel Processing Technology, vol. 98, pp. 23–29, 2012. View at Publisher · View at Google Scholar · View at Scopus
  28. M. S. M. Ali, C. C. Yun, A. L. T. Chor, R. N. Z. R. A. Rahman, M. Basri, and A. B. Salleh, “Purification and characterisation of an F16L mutant of a thermostable lipase,” Protein Journal, vol. 31, no. 3, pp. 229–237, 2012. View at Publisher · View at Google Scholar · View at Scopus
  29. J.-J. Shangguan, Y.-Q. Liu, F.-J. Wang et al., “Expression and characterization of a novel lipase from aspergillus fumigatus with high specific activity,” Applied Biochemistry and Biotechnology, vol. 165, no. 3-4, pp. 949–962, 2011. View at Publisher · View at Google Scholar · View at Scopus
  30. V. M. G. Lima, N. Krieger, D. A. Mitchell, and J. D. Fontana, “Activity and stability of a crude lipase from Penicillium aurantiogriseum in aqueous media and organic solvents,” Biochemical Engineering Journal, vol. 18, no. 1, pp. 65–71, 2004. View at Publisher · View at Google Scholar · View at Scopus
  31. A. Hiol, M. D. Jonzo, N. Rugani, D. Druet, L. Sarda, and L. C. Comeau, “Purification and characterization of an extracellular lipase from a thermophilic Rhizopus oryzae strain isolated from palm fruit,” Enzyme and Microbial Technology, vol. 26, no. 5-6, pp. 421–430, 2000. View at Publisher · View at Google Scholar · View at Scopus
  32. C. M. F. Soares, H. F. de Castro, F. F. de Moraes, and G. M. Zanin, “Characterization and utilization of Candida rugosa lipase immobilized on controlled pore silica,” Applied Biochemistry and Biotechnology, vol. 77–79, pp. 745–757, 1999. View at Google Scholar · View at Scopus
  33. S. Hama, S. Tamalampudi, T. Fukumizu et al., “Lipase localization in Rhizopus oryzae cells immobilized within biomass support particles for use as whole-cell biocatalysts in biodiesel-fuel production,” Journal of Bioscience and Bioengineering, vol. 101, no. 4, pp. 328–333, 2006. View at Publisher · View at Google Scholar · View at Scopus
  34. A. M. Klibanov, “Improving enzymes by using them in organic solvents,” Nature, vol. 409, no. 6817, pp. 241–246, 2001. View at Publisher · View at Google Scholar · View at Scopus
  35. A. Jayaprakash and P. Ebenezer, “Purification and characterization of Aspergillus japonicus lipase from a pig fat production medium,” Journal of Academia and Industrial Research, vol. 1, no. 1, pp. 1–7, 2012. View at Google Scholar
  36. D. Pokorny, A. Cimerman, and W. Steiner, “Aspergillus niger lipases: induction, isolation and characterization of two lipases from a MZKI A116 strain,” Journal of Molecular Catalysis B: Enzymatic, vol. 2, no. 4-5, pp. 215–222, 1997. View at Publisher · View at Google Scholar · View at Scopus
  37. N. C. Mhetras, K. B. Bastawde, and D. V. Gokhale, “Purification and characterization of acidic lipase from Aspergillus niger NCIM 1207,” Bioresource Technology, vol. 100, no. 3, pp. 1486–1490, 2009. View at Publisher · View at Google Scholar · View at Scopus
  38. M. de Fátima Silva Lopes, A. L. Leitão, M. Regalla, J. J. F. Marques, M. J. T. Carrondo, and M. T. B. Crespo, “Characterization of a highly thermostable extracellular lipase from Lactobacillus plantarum,” International Journal of Food Microbiology, vol. 76, no. 1-2, pp. 107–115, 2002. View at Publisher · View at Google Scholar · View at Scopus
  39. N. R. Kamini, J. G. S. Mala, and R. Puvanakrishnan, “Lipase production from Aspergillus niger by solid-state fermentation using gingelly oil cake,” Process Biochemistry, vol. 33, no. 5, pp. 505–511, 1998. View at Publisher · View at Google Scholar · View at Scopus
  40. A. Tsang, G. Butler, J. Powlowski, E. A. Panisko, and S. E. Baker, “Analytical and computational approaches to define the Aspergillus niger secretome,” Fungal Genetics and Biology : FG & B, vol. 46, pp. S153–S160, 2009. View at Publisher · View at Google Scholar · View at Scopus
  41. Z.-Y. Shu, J.-K. Yang, and Y.-J. Yan, “Purification and characterization of a lipase from Aspergillus niger F044,” Sheng Wu Gong Cheng Xue Bao, vol. 23, no. 1, pp. 96–101, 2007. View at Publisher · View at Google Scholar · View at Scopus
  42. S. M. Basheer, S. Chellappan, P. S. Beena, R. K. Sukumaran, K. K. Elyas, and M. Chandrasekaran, “Lipase from marine Aspergillus awamori BTMFW032: production, partial purification and application in oil effluent treatment,” New Biotechnology, vol. 28, no. 6, pp. 627–638, 2011. View at Publisher · View at Google Scholar · View at Scopus
  43. H. S. Hamdy and M. A. Abo-Tahon, “Extracellular lipase of Aspergillus terreus var. africanus (CBS 130.55): production, purification and characterisation,” Annals of Microbiology, vol. 62, no. 4, pp. 1723–1736, 2012. View at Publisher · View at Google Scholar · View at Scopus
  44. C. M. Romero, L. M. Pera, F. Loto, and M. D. Baigori, “Specific enzyme-catalyzed hydrolysis and synthesis in aqueous and organic medium using biocatalysts with lipase activity from Aspergillus niger MYA 135,” Catalysis Letters, vol. 142, no. 11, pp. 1361–1368, 2012. View at Publisher · View at Google Scholar · View at Scopus
  45. J. C. Mateos Diaz, J. A. Rodríguez, S. Roussos et al., “Lipase from the thermotolerant fungus Rhizopus homothallicus is more thermostable when produced using solid state fermentation than liquid fermentation procedures,” Enzyme and Microbial Technology, vol. 39, no. 5, pp. 1042–1050, 2006. View at Publisher · View at Google Scholar · View at Scopus
  46. M. A. F. Costa and R. M. Peralta, “Production of lipase by soil fungi and partial characterization of lipase from a selected strain (Penicillium wortmanii),” Journal of Basic Microbiology, vol. 39, no. 1, pp. 11–15, 1999. View at Publisher · View at Google Scholar
  47. A. Ibrik, H. Chahinian, N. Rugani, L. Sarda, and L.-C. Comeau, “Biochemical and structural characterization of triacylglycerol lipase from Penicillium cyclopium,” Lipids, vol. 33, no. 4, pp. 377–384, 1998. View at Publisher · View at Google Scholar · View at Scopus
  48. E. Rigo, J. L. Ninow, S. M. Tsai et al., “Preliminary characterization of novel extra-cellular lipase from Penicillium crustosum under solid-state fermentation and its potential application for triglycerides hydrolysis,” Food and Bioprocess Technology, vol. 5, no. 5, pp. 1592–1600, 2012. View at Publisher · View at Google Scholar · View at Scopus
  49. L. L. Zhao, J. H. Xu, J. Zhao, J. Pan, and Z. L. Wang, “Biochemical properties and potential applications of an organic solvent-tolerant lipase isolated from Serratia marcescens ECU1010,” Process Biochemistry, vol. 43, no. 6, pp. 626–633, 2008. View at Publisher · View at Google Scholar · View at Scopus
  50. P. Sarkar, S. Yamasaki, S. Basak, A. Bera, and P. K. Bag, “Purification and characterization of a new alkali-thermostable lipase from Staphylococcus aureus isolated from Arachis hypogaea rhizosphere,” Process Biochemistry, vol. 47, no. 5, pp. 858–866, 2012. View at Publisher · View at Google Scholar · View at Scopus
  51. J. Dong, P. Deng, and J. Xu, “Study of the effects of Cr ions on Yb in Cr,Yb:YAG crystal,” Optics Communications, vol. 170, no. 4–6, pp. 255–258, 1999. View at Publisher · View at Google Scholar · View at Scopus
  52. J. M. Palomo, G. Fernández-Lorente, J. M. Guisán, and R. Fernández-Lafuente, “Modulation of immobilized lipase enantioselectivity via chemical amination,” Advanced Synthesis and Catalysis, vol. 349, no. 7, pp. 1119–1127, 2007. View at Publisher · View at Google Scholar · View at Scopus
  53. G. Fernandez-Lorente, J. M. Palomo, Z. Cabrera, R. Fernandez-Lafuente, and J. M. Guisán, “Improved catalytic properties of immobilized lipases by the presence of very low concentrations of detergents in the reaction medium,” Biotechnology and Bioengineering, vol. 97, no. 2, pp. 242–250, 2007. View at Publisher · View at Google Scholar · View at Scopus
  54. B. S. Demir and S. S. Tükel, “Purification and characterization of lipase from Spirulina platensis,” Journal of Molecular Catalysis B: Enzymatic, vol. 64, no. 3-4, pp. 123–128, 2010. View at Publisher · View at Google Scholar · View at Scopus
  55. G. Fernández-Lorente, J. M. Palomo, M. Fuentes, C. Mateo, J. M. Guisán, and R. Fernández-Lafuente, “Self-assembly of Pseudomonas fluorescens lipase into bimolecular aggregates dramatically affects functional properties,” Biotechnology and Bioengineering, vol. 82, no. 2, pp. 232–237, 2003. View at Publisher · View at Google Scholar · View at Scopus
  56. C. Yang, F. Wang, D. Lan, C. Whiteley, B. Yang, and Y. Wang, “Effects of organic solvents on activity and conformation of recombinant Candida antarctica lipase A produced by Pichia pastoris,” Process Biochemistry, vol. 47, no. 3, pp. 533–537, 2012. View at Publisher · View at Google Scholar · View at Scopus
  57. E. H. Ahmed, T. Raghavendra, and D. Madamwar, “An alkaline lipase from organic solvent tolerant Acinetobacter sp. EH28: application for ethyl caprylate synthesis,” Bioresource Technology, vol. 101, no. 10, pp. 3628–3634, 2010. View at Publisher · View at Google Scholar · View at Scopus
  58. L. Daoud, J. Kamoun, M. B. Ali et al., “Purification and biochemical characterization of a halotolerant Staphylococcus sp. extracellular lipase,” International Journal of Biological Macromolecules, vol. 57, pp. 232–237, 2013. View at Publisher · View at Google Scholar · View at Scopus
  59. D. S. Dheeman, S. Antony-Babu, J. M. Frías, and G. T. M. Henehan, “Purification and characterization of an extracellular lipase from a novel strain Penicillium sp. DS-39 (DSM 23773),” Journal of Molecular Catalysis B: Enzymatic, vol. 72, no. 3-4, pp. 256–262, 2011. View at Publisher · View at Google Scholar · View at Scopus
  60. X. Li, P. Qian, S.-G. Wu, and H.-Y. Yu, “Characterization of an organic solvent-tolerant lipase from Idiomarina sp. W33 and its application for biodiesel production using Jatropha oil,” Extremophiles, vol. 18, no. 1, pp. 171–178, 2014. View at Publisher · View at Google Scholar · View at Scopus
  61. J.-L. Xia, B. Huang, Z.-Y. Nie, and W. Wang, “Production and characterization of alkaline extracellular lipase from newly isolated strain Aspergillus awamori HB-03,” Journal of Central South University of Technology, vol. 18, no. 5, pp. 1425–1433, 2011. View at Publisher · View at Google Scholar · View at Scopus
  62. I. Mayordomo, F. Randez-Gil, and J. A. Prieto, “Isolation, purification, and characterization of a cold-active lipase from Aspergillus nidulans,” Journal of Agricultural and Food Chemistry, vol. 48, no. 1, pp. 105–109, 2000. View at Publisher · View at Google Scholar · View at Scopus
  63. N. R. Kamini, T. Fujii, T. Kurosu, and H. Iefuji, “Production, purification and characterization of an extracellular lipase from the yeast, Cryptococcus sp. S-2,” Process Biochemistry, vol. 36, no. 4, pp. 317–324, 2000. View at Publisher · View at Google Scholar · View at Scopus
  64. K. Yamamoto and N. Fujiwara, “The hydrolysis of castor oil using a lipase from Pseudomonas sp. f-B-24: positional and substrate specificity of the enzyme and optimum reaction conditions,” Bioscience, Biotechnology and Biochemistry, vol. 59, no. 7, pp. 1262–1266, 1995. View at Publisher · View at Google Scholar · View at Scopus