Appl Biochem Biotechnol DOI 10.1007/s12010-014-1159-9
Production of Thermostable Lipase by Thermomyces lanuginosus on Solid-State Fermentation: Selective Hydrolysis of Sardine Oil Nayeli Ávila-Cisneros & Susana Velasco-Lozano & Sergio Huerta-Ochoa & Jesús Córdova-López & Miquel Gimeno & Ernesto Favela-Torres
Received: 22 April 2014 / Accepted: 15 August 2014 # Springer Science+Business Media New York 2014
Abstract A naturally immobilized biocatalyst with lipase activity was produced by Thermomyces lanuginosus on solid-state fermentation with perlite as inert support. Maxima lipase activities (22 and 120 U/g of dry matter, using p-nitrophenyl octanoate and trioctanoine, respectively, as substrates) were obtained after 72 h of solid culture, remaining nearly constant up to 120 h. Maxima lipase activity was found at 60 to 85 °C and pH 10. The biocatalyst was stable at 60 °C for at least 4 h of incubation and a pH from 7 to 10. Energy values of activation and deactivation of lipase were of 26 and 6.7 kJ/mol, respectively. The biocatalyst shows high selectivity for the release of the omega-3 polyunsaturated fatty acids, eicosapentaenoic (EPA) and docosahexaenoic acids (DHA), during the hydrolysis of sardine oil. The EPA/DHA ratio (16:6) released by this biocatalyst was superior to that obtained with the commercial preparations of T. lanuginosus. Keywords Solid-state fermentation . Thermomyces lanuginosus . Thermostable lipases . Sardine oil hydrolysis
N. Ávila-Cisneros : S. Velasco-Lozano : S. Huerta-Ochoa : E. Favela-Torres (*) Universidad Autónoma Metropolitana—Unidad Iztapalapa, Av. San Rafael Atlixco 186, Col. Vicentina, Iztapalapa, Mexico 09340 D.F., Mexico e-mail: [email protected] J. Córdova-López Departamento de Química, Universidad de Guadalajara, Guadalajara, Mexico M. Gimeno Facultad de Química, Departamento de Alimentos y Biotecnología, Universidad Nacional Autónoma de México, Ciudad Universitaria, Mexico, D.F. 04510, Mexico Present Address: E. Favela-Torres Departamento de Biotecnología, Universidad Autónoma Metropolitana—Unidad Iztapalapa, Av. San Rafael Atlixco 186, Col. Vicentina, Iztapalapa, Mexico 09340 D.F., Mexico
Appl Biochem Biotechnol
Introduction Supplementation of human diet with the long-chain omega-3 polyunsaturated free fatty acids (PUFAs), such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), has health benefits owing to the reduction of cardiovascular risks [1]. In this regard, fish oil is the main source of EPA and DHA. In most lipids of fish species, the EPA/DHA ratio is lower than 1; however, the requirements for commercial supplements, this ratio is higher than 1 (up to 3.1) [2]. Therefore, chemical or enzymatic separation of PUFAs is required to formulate well-balanced supplements. Nonetheless, separation of EPA and DHA is rather complex owing to their physical and structural similarities. Physicochemical methods under supercritical conditions proved successful [3]; however, enzymatic routes offer mild and nontoxic alternative procedures [4]. Additionally, lipases might exhibit high regio- and acyl-selectivity toward some fatty acids of triacylglycerides [5]. Studies with different lipases and substrates showed that specificity toward EPA and DHA strongly depends on both regio- and acyl-selectivity of lipases and the nature of substrates [6]. Immobilization of enzymes confers improved enzyme catalytic properties as enhanced stability, activity, and selectivity as demonstrated by several groups [7–10]. Regarding PUFA concentration, different immobilization strategies have allowed EPA/DHA ratios up to 34 and 12 after fish oil hydrolysis with commercial immobilized lipases from Rhizomucor miehei and Thermomyces lanuginosus, respectively [4]. EPA/DHA ratio near to 3 was obtained after 70 % hydrolysis of anchovy oil with the lipase of T. lanuginosus (TLL) [11]. It is worth noting that aspects such as type of lipases, immobilization protocol, and reaction conditions affect the activity and selectivity of the hydrolytic reaction of fats and oils. TLL is an important biocatalyst for fat and oil modifications, enantio- and regio-selective reactions and resolution of racemic mixtures, among others [12]. T. lanuginosus is a thermophile fungus producing extracellular thermostable enzymes, such as xylanases [13], amylases, glycosidases [14], and lipases [15, 16]. Lipases from T. lanuginosus are mainly produced at industrial scale with well-known recombinant strains of Aspergillus niger, but studies leading to lipase production by T. lanuginosus, in either submerged or solid-state cultures, are scarce [15, 17]. The production of lipases from T. lanuginosus by solid-state fermentation (SSF) is advantageous in terms of higher production and productivity of enzymes, reduced catabolite repression, better culture oxygenation, and enzymes with improved catalytic characteristics (higher thermoactivity and stability) [18, 19]. Additionally, the low transfer rate of metabolic heat in SSF [20] might enhance growth and enzyme production of thermophile fungi, such as T. lanuginosus. Moreover, the use of SSF also allows obtaining a solid biocatalyst with “naturally immobilized” enzymes, preventing unit operations such as extraction, concentration, purification, and further immobilization of enzymes [21]. Relevant aspects related to SSF processes have broadly been reviewed [22, 23]; however, to our knowledge, there are no reports on the use of inert supports for the production of biocatalysts with lipase activity from T. lanuginosus. In this study, the production and characterization of a thermostable lipase from T. lanuginosus by solid-state fermentation were achieved. The fermented solids (biocatalyst) were employed for the selective hydrolysis of sardine oil.
Methods Fungal Strain T. lanuginosus SS-2 isolated from sugar cane bagasse (Gen Bank ID: JQ639283) was supplied by Dr. Jesus Cordova at the University of Guadalajara (Mexico). This strain was lyophilized for storage.
Appl Biochem Biotechnol
Pre-inoculum Preparation Petri dishes containing 30 mL of potato dextrose agar (PDA) covered with a cellophane membrane (Cellophane No. 5, obtained from Central Supplies in Mexico City) were inoculated with a piece of mycelium (0.5×0.5 cm) obtained from a previous culture on PDA at 45 °C for 7 days. PDA and cellophane membranes were separately sterilized at 121 °C for 15 min. Culture Media Modified Pontecorvo medium with the following composition was used for inoculum production (in g/L): (NH4)2SO4, 4.63; urea, 3.16; KH2PO4, 14; K2HPO4, 2.4; KCl, 0.8; MgSO4 · 7H2O, 0.8; yeast extract, 0.8; glucose, 28.8; olive oil, 7.2; polyvinyl alcohol, 2.1; and 1.2 mL of trace element solution containing (in mg/L) 100 Na2B4O7 · 10H2O, 50 MnCl2 · 4H2O, 50 Na2MoO4 · 2H2O, 250 CuSO4 · 5H2O, 85 FeCl3 · 6H2O, and 100 ZnSO4 · 7H2O. Culture medium for lipase production by SSF was 3-fold concentrated. Initial pH was adjusted to 6.5. Inoculum Preparation Two pieces of mycelia (0.5×0.5 cm) over cellophane membrane, obtained from a preinoculum, were cut and transferred to a sterile tube containing 5 mL of sterile culture medium and 4 g of glass bead (3 mm, Marienfeld Laboratory Glassware). Mycelium was gently disaggregated by manual agitation for 2 min. Two milliliters of the obtained suspension, free of glass bead, was added to a 125-mL Erlenmeyer flask containing 28 mL of culture medium. Flasks were incubated at 40 °C in an orbital shaker (150 rpm) for 72 h. The obtained culture was used to inoculate the culture medium for SSF. Production of Lipases by SSF Perlite was used as an inert support. It was previously washed with distilled water, dried, and sieved (mesh 18/20). Support was impregnated with inoculated (10 % v/v) culture medium (65 mL of inoculated medium and 35 g of perlite) to obtain an initial moisture content of 65 % (w/v). Glass columns for SSF (2.5-cm diameter×20 cm long) [24] were filled with 8 g of the impregnated solid medium and incubated at 45 °C for 120 h, supplying water-saturated air at a flow rate of 50 mL/min. Two columns were sampled at different time intervals for lipase, pH, and moisture content assays. CO2 production was monitored online by connecting the outlet air stream to a gas analyzer. CO2 production rate, specific CO2 production rate (μCO2), and lag phase were estimated based on the CO2 concentration in the exhaust gas stream [25]. Sample Treatment Lipase activity was assayed using enzymatic extracts or dry biocatalysts [21]. The enzymatic extracts were obtained from 1 g of dried fermented material suspended in 10 mL of distilled water, agitated in vortex for 2 min, and centrifuged at 10,000 rpm for 3 min. Supernatants were kept at 4 °C for further assays (lipase activity, pH, and glucose). The dry biocatalysts were obtained by blowing air (at 10 L/min) into a glass column containing the fermented solid to a final moisture content below 1 % w/v. The dry fermented solids were milled with a mortar to obtain a fine powder that was stored at room temperature in the dark. This dry biocatalyst maintained its full activity for at least 3 months. Moisture content was determined by using a thermobalance (Ohaus Model MB45, USA).
Appl Biochem Biotechnol
Enzyme Immobilization For the hydrolysis studies of sardine oil, lipases from the dry biocatalyst were extracted and immobilized as follows: In order to get similar enzymatic activity for the preparation of the biocatalyst and the octyl-agarose-immobilized enzyme, 1 g of dry biocatalyst and 10 mL of cold (4 °C) distilled water were mixed for 2 min. Supernatants were collected by centrifugation at 8,000 rpm for 10 min. Ten milliliters of supernatant were added to 1 g of octyl-agarose and maintained under mild agitation at 4 °C until no activity was detected in the filtered supernatant (1 h). In order to evaluate the loss of activity due to the immobilization procedure, the activity of the enzyme solution without support was monitored at the same conditions. Finally, the immobilized enzyme was washed three times with 10 mL of phosphate buffer (25 mM, pH 7) and stored at 4 °C. Enzymatic Activity Assays Spectrophotometric Assay The substrate solution contained one volume of 10 mM pnitrophenyl octanoate (p-NPO) (dissolved in 2-propanol) and nine volumes of 100 mM of Tris–HCl (pH 8) and 0.25 % (w/v) of polyvinyl alcohol (PVA) [19]. The enzymatic reaction was started by adding 50 μL of enzymatic extract to 100 μL of substrate solution. Absorbance was monitored at 405 nm using a multiwell-microplate reader (Biotek Elx808, USA) at 30 °C for 5 min. The reaction rate was calculated from the slope of a standard curve of absorbance versus time, using a molar extinction coefficient of 7,859/cm M) for p-nitrophenol at the assay conditions (pH 8, 30 °C). For all assays, the hydrolysis rate of a control (buffer instead of enzymatic extract) was recorded and subtracted. One unit of enzyme activity (U) was defined as the amount of enzyme releasing 1 μmol of p-nitrophenol per minute. Titrimetric Assay The substrate emulsion was prepared as follows: 10 mL of trioctanoin and 90 mL of 10 % (w/v) gum arabic at pH 7 were emulsified with a mechanical homogenizer (Oster blender with two speeds, Model 2523-13 440-20, Mexico) for 10 min. The trioctanoin emulsion (6 mL) was mixed with 24 mL of 0.3 mM Tris–HCl (pH 7) containing 150 mM NaCl and 2 mM CaCl2. The mixture was pre-incubated at the selected temperature, and the reaction was started by adding the enzymatic extract (adequately diluted) or the biocatalyst. Initial pH was set at 7, and the released fatty acids were continuously titrated with 0.1 M NaOH for 5 min, using an automatic titrator (Mettler DL21 Titrator, Switzerland). Each assay was performed by triplicate. One unit of enzymatic activity (U) was defined as the amount of enzyme releasing 1 μmol of fatty acid per minute [19]. Possible proteases contained in the enzymatic extracts were inhibited using a cocktail specific for fungal proteases (P8215; Sigma); for that, 10 μL of the protease inhibitor (2 mM) was added to 50 μL of enzymatic extract, and the lipase activity was spectrophotometrically estimated using p-NPO at 30 °C and pH 8. Effect of Temperature on Lipase Activity and Stability The effect of temperature on lipase activity was studied using the titrimetric assay. Trioctanoin emulsions were pre-incubated at temperatures from 30 to 85 °C. To start the enzymatic reaction, 50 mg of biocatalyst was added and continuously titrated with 0.1 M NaOH for 5 min under mild agitation. Lipase activity was assayed by triplicate, and the obtained values were used to estimate activation (Ea) and deactivation (Ed) energy, according to the Arrhenius equation. The lipase stability was evaluated using the spectrophotometric assay with p-NPO as
Appl Biochem Biotechnol
substrate. Enzymatic extracts were pre-incubated at temperatures from 30 to 80 °C for 4 h, taking samples every 30 min and assaying the residual lipase activity at 30 °C. Effect of pH on Lipase Activity and Stability The effect of pH on the lipase activity at 30 °C and stability at 4 °C was evaluated spectrophotometrically at 348 nm. Enzymatic extracts with 100 mM p-NPO as substrate (for activity assays) or without p-NPO (for stability assays) were incubated at different pH values. For that, the following buffers were used (at 0.1 M): citrate–phosphate, pH 3 to 6, Tris–HCl, pH 7 to 9, NaHCO3–NaOH, pH 10–11, and KCl- NaOH, pH 12. For enzymatic stability, residual activity was spectrophotometrically assayed at pH 8 and 30 °C for 5 min. Hydrolysis of Sardine Oil Sardine oil was hydrolyzed following the methodology proposed by Fernández-Lorente et al. [4]. One gram of dry biocatalyst or octyl-immobilized biocatalyst was added to a 15-mL flask containing 10 mL of a biphasic mixture of 0.1 mL of sardine oil in 4.9 mL of cyclohexane/ 5 mL of aqueous buffer solution (10-mM phosphate buffer for pH 6 or 10 mM Tris–HCl for pH 8). The flasks were covered and maintained under gentle magnetic stirring at different temperatures for different time intervals. One unit was defined as the amount of enzyme required to release 1 μmol of EPA and/or DHA per minute under the above conditions. Experiments were carried out by duplicate with coefficients of variation lower than 5 %. Analysis of EPA and DHA by HPLC-UV Samples were taken at different time intervals and used to determine the released PUFAs; for that, 50 μL of sample was dissolved in 400 μL of acetonitrile and analyzed by HPLC (Spectra Physic HPLC-UV) with an UV detector (SP 100–UV detector SP 8450) and equipped with a Kromasil C8 (15×0.4 mm) column. Samples were eluted with acetonitrile/water/acetic acid (70:30:0.1) at pH 3 as mobile phase at an isocratic flow rate of 1 mL/min. PUFAs were detected at 215 nm. Retention times were 10.3 min for EPA and 13.2 min for DHA.
Results and Discussions Kinetics of CO2 Production of T. lanuginosus Cultured on SSF Kinetics of CO2 production and CO2 production rate are shown in Fig. 1. The lag phase of 13.4 h was similar to that reported for submerged cultures of T. lanuginosus [26] and a recombinant strain of A. niger, expressing a lipase-encoding gene from T. lanuginosus [27]. Maximum CO2 production rate (3.7 mg CO2/g initial dry matter (idm) h) was reached after 31 h of culture. At the end of culture (120 h), 116 mg CO2/g idm was produced (Fig. 1a). It is worth to mention that due to the difficulties to estimate the microbial growth on solid cultures, kinetic parameters of T. lanuginosus on SSF have not been reported. The specific CO2 production rate obtained in this study (0.11/h) is slightly lower than that reported for the recombinant strain of A. niger, expressing the lipase of T. lanuginosus [27]. In related work, Gomes et al. [26] found specific growth rates from 0.1 to 0.31/h for T. lanuginosus growing in submerged cultures. On the other hand, high specific growth rate values (0.89 and 0.46/h) were observed during the biphasic growth of T. lanuginosus in rich YPS medium [28].
Appl Biochem Biotechnol
Lipase Production by T. lanuginosus Cultured on SSF
120
4 a) 3
80 2 40 1
0
Production of CO2 (mg/g idm)
CO2 production rate (mg/g idm h)
Although glucose is a well-recognized catabolic repressor, previous studies report low catabolite repression during the production of pectinases [29] and tannases [30] by A. niger cultured on SSF. In this study, lipase production started once glucose was completely consumed after 24 h of culture (results not shown), reaching a maximum value of 22±2.2 U/g dm at 48 h of culture (Fig. 1b) and remaining nearly constant (19.5±1.8 U/g dm) until the end of culture (120 h). The use of the Luedecking–Piret equation [31] allowed showing that lipase production was directly associated to CO2 production (α=0.244 U/mg) with a nonassociated level (β=−9× 10−4 U/mg h) nearly negligible at the end of culture (Table 1). The high absolute value of η (−30) suggested a high degree of correlation between lipase production and CO2 production [18].
0
0
24
48
72
96
120
Time (h)
Lipase Activity (U/g dm)
30
b)
25 20 15 10 5 0 0
24
48
72 Time (h)
96
120
Fig. 1 Kinetics of CO2 production (mg/g idm—points) and CO2 production rate (mg/g idm h—continuous line) (a) and lipase production (b) of T. lanuginosus cultured on SSF at 40 °C, using perlite as inert support impregnated with a culture medium
Appl Biochem Biotechnol
Table 1 Kinetic parameters of T. lanuginosus growth cultured on SSF
α production of enzymatic activity associated to the production of CO2, η ratio αμ/β, β rate of enzymatic activity not associated to the production of CO2
Parameter
Value
CO2max (mg/g idm)
116.4±10.7
CO2 (mg/g idm h) μ (1/h)
3.5±0.4 0.11±0.01
Lag phase (h)
13.4±0.5
α (U/mg CO2)
0.244
β (U/mg CO2 h)
−0.0009
η
−30
Additionally, the low β value suggested the absence of proteases at the end of culture. This was corroborated assaying lipase activity in the presence of a proteases inhibitor; finding that activity remained constant with or without the inhibitor. Catalytic characteristics of the biocatalysts produced in this work were carried out using either enzymatic extracts or dry biocatalysts. For that, SSF samples were obtained at the same physiological state, namely, once the maximum CO2 production rate was achieved (around 44 h of culture, see Fig. 2a). The physiological state of culture was monitored by online analysis of CO2 in the exhaust air phase. Lipase activity was titrimetrically assayed with trioctanoin for both the enzymatic extract (liquid) and the biocatalyst (solid) samples, obtaining 58±1 and 63±2 U/g dm, respectively. Lipase activity was also assayed with the spectrophotometric method with p-nitrophenyl octanoate as substrate. Twenty times higher activity was obtained with the titrimetric method. Likewise, measurement of lipase activity with olive oil as substrate yielded enzymatic titers up to 13.5 higher than that obtained with p-nitrophenyl palmitate as substrate [24]. Effect of Temperature on Lipase Activity The effect of temperature on lipase activity of T. lanuginosus was evaluated by incubating the enzymatic extracts from 30 to 85 °C. Lipase activity increased up to 2.0-fold from 30 to 60 °C and at higher incubation temperatures (from 60 to 85 °C); the lipase activity remained nearly constant (Fig. 2a). To our knowledge, 85 °C has been the highest record reported for lipases produced by filamentous fungi (Table 2). Similar profile of lipase activity as a function of temperature was obtained using the solid biocatalyst and the titrimetric assay (results not shown). Lipase activity at 85 °C is higher than the reported (80 °C) for the well-known immobilized lipase B of Candida antarctica [32]. High temperatures have also been reported (70 °C) for xylanases and (70 °C) for phytases from T. lanuginosus [33]. The calculated activation energy (Ea=27 kJ/mol) for the lipase produced in this work was similar to those reported for the lipases from Candida rugosa (30 to 40 kJ/mol) [34] and from Rhizopus homothallicus (30 kJ/mol) [19] (Table 3). However, higher Ea value was reported (124.8 kJ/mol) for xylanases from T. lanuginosus [36]. The calculated deactivation energy (Ed=6.7 kJ/mol) for the lipase produced in this work was lower than those reported for a commercial lipase from T. lanuginosus (26.5 kJ/mol) [37] and for xylanases produced by wild and mutants strains of T. lanuginosus (98.6 and 71.8 kJ/mol, respectively) [36]. The high Ea/ Ed ratio obtained herein (4.0) gives evidence about the potential application of the biocatalyst at high temperatures of reaction [38, 39]; for instance, the acidolysis reactions for the production of structured lipids are carried out from 45 to 50 °C [40].
Appl Biochem Biotechnol
Lipase activity (U/g dm)
150
a)
120 90 60 30 0 30
Relative activity (%)
120
40
50 60 70 Temperature ( C)
80
90
b)
90
60
30
0 3
4
5
6
7
8
9
10
11
12
pH Fig. 2 Effect of temperature (a) and pH (b) on lipase activity of T. lanuginosus produced on SSF. Error bars represent the standard deviation of three independent assays. Activity was titrimetrically assayed with trioctanoin as substrate
Effect of Temperature on Lipase Stability The thermostability studies carried out with the enzymatic extracts of T. lanuginosus showed that lipase activity was enhanced from 30 to 50 °C and maintained constant at 60 °C, after an incubation time of 4 h. At 70 °C, enzyme activity decreased 50 % after being incubated in 0.5 h. An increase of activity up to 30 % was observed from 30 to 60 °C after 2 h of incubation. Some authors have discussed the high thermal stability and improvement in
Appl Biochem Biotechnol Table 2 Effect of temperature on activity and stability of lipases produced by solid-state fermentation (SSF) and submerged fermentation (SmF) for different fungi strains Fungi
Topt (°C)
T* (°C)
T. lanuginosus
80
70
0.5
80
80
0.25
30
50
0.4
nr
SmF
40
50
0.7
nr
SSF
37
50
0.7
1.0
cp
50
50
1.1
0.6
cp
Penicillium sp.
37
65
1.5
1.9
SSF
Humicola lanuginosa
42 45
25 70
0.3 ≈2
Production of Thermostable Lipase by Thermomyces lanuginosus on Solid-State Fermentation: Selective Hydrolysis of Sardine Oil Nayeli Ávila-Cisneros & Susana Velasco-Lozano & Sergio Huerta-Ochoa & Jesús Córdova-López & Miquel Gimeno & Ernesto Favela-Torres
Received: 22 April 2014 / Accepted: 15 August 2014 # Springer Science+Business Media New York 2014
Abstract A naturally immobilized biocatalyst with lipase activity was produced by Thermomyces lanuginosus on solid-state fermentation with perlite as inert support. Maxima lipase activities (22 and 120 U/g of dry matter, using p-nitrophenyl octanoate and trioctanoine, respectively, as substrates) were obtained after 72 h of solid culture, remaining nearly constant up to 120 h. Maxima lipase activity was found at 60 to 85 °C and pH 10. The biocatalyst was stable at 60 °C for at least 4 h of incubation and a pH from 7 to 10. Energy values of activation and deactivation of lipase were of 26 and 6.7 kJ/mol, respectively. The biocatalyst shows high selectivity for the release of the omega-3 polyunsaturated fatty acids, eicosapentaenoic (EPA) and docosahexaenoic acids (DHA), during the hydrolysis of sardine oil. The EPA/DHA ratio (16:6) released by this biocatalyst was superior to that obtained with the commercial preparations of T. lanuginosus. Keywords Solid-state fermentation . Thermomyces lanuginosus . Thermostable lipases . Sardine oil hydrolysis
N. Ávila-Cisneros : S. Velasco-Lozano : S. Huerta-Ochoa : E. Favela-Torres (*) Universidad Autónoma Metropolitana—Unidad Iztapalapa, Av. San Rafael Atlixco 186, Col. Vicentina, Iztapalapa, Mexico 09340 D.F., Mexico e-mail: [email protected] J. Córdova-López Departamento de Química, Universidad de Guadalajara, Guadalajara, Mexico M. Gimeno Facultad de Química, Departamento de Alimentos y Biotecnología, Universidad Nacional Autónoma de México, Ciudad Universitaria, Mexico, D.F. 04510, Mexico Present Address: E. Favela-Torres Departamento de Biotecnología, Universidad Autónoma Metropolitana—Unidad Iztapalapa, Av. San Rafael Atlixco 186, Col. Vicentina, Iztapalapa, Mexico 09340 D.F., Mexico
Appl Biochem Biotechnol
Introduction Supplementation of human diet with the long-chain omega-3 polyunsaturated free fatty acids (PUFAs), such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), has health benefits owing to the reduction of cardiovascular risks [1]. In this regard, fish oil is the main source of EPA and DHA. In most lipids of fish species, the EPA/DHA ratio is lower than 1; however, the requirements for commercial supplements, this ratio is higher than 1 (up to 3.1) [2]. Therefore, chemical or enzymatic separation of PUFAs is required to formulate well-balanced supplements. Nonetheless, separation of EPA and DHA is rather complex owing to their physical and structural similarities. Physicochemical methods under supercritical conditions proved successful [3]; however, enzymatic routes offer mild and nontoxic alternative procedures [4]. Additionally, lipases might exhibit high regio- and acyl-selectivity toward some fatty acids of triacylglycerides [5]. Studies with different lipases and substrates showed that specificity toward EPA and DHA strongly depends on both regio- and acyl-selectivity of lipases and the nature of substrates [6]. Immobilization of enzymes confers improved enzyme catalytic properties as enhanced stability, activity, and selectivity as demonstrated by several groups [7–10]. Regarding PUFA concentration, different immobilization strategies have allowed EPA/DHA ratios up to 34 and 12 after fish oil hydrolysis with commercial immobilized lipases from Rhizomucor miehei and Thermomyces lanuginosus, respectively [4]. EPA/DHA ratio near to 3 was obtained after 70 % hydrolysis of anchovy oil with the lipase of T. lanuginosus (TLL) [11]. It is worth noting that aspects such as type of lipases, immobilization protocol, and reaction conditions affect the activity and selectivity of the hydrolytic reaction of fats and oils. TLL is an important biocatalyst for fat and oil modifications, enantio- and regio-selective reactions and resolution of racemic mixtures, among others [12]. T. lanuginosus is a thermophile fungus producing extracellular thermostable enzymes, such as xylanases [13], amylases, glycosidases [14], and lipases [15, 16]. Lipases from T. lanuginosus are mainly produced at industrial scale with well-known recombinant strains of Aspergillus niger, but studies leading to lipase production by T. lanuginosus, in either submerged or solid-state cultures, are scarce [15, 17]. The production of lipases from T. lanuginosus by solid-state fermentation (SSF) is advantageous in terms of higher production and productivity of enzymes, reduced catabolite repression, better culture oxygenation, and enzymes with improved catalytic characteristics (higher thermoactivity and stability) [18, 19]. Additionally, the low transfer rate of metabolic heat in SSF [20] might enhance growth and enzyme production of thermophile fungi, such as T. lanuginosus. Moreover, the use of SSF also allows obtaining a solid biocatalyst with “naturally immobilized” enzymes, preventing unit operations such as extraction, concentration, purification, and further immobilization of enzymes [21]. Relevant aspects related to SSF processes have broadly been reviewed [22, 23]; however, to our knowledge, there are no reports on the use of inert supports for the production of biocatalysts with lipase activity from T. lanuginosus. In this study, the production and characterization of a thermostable lipase from T. lanuginosus by solid-state fermentation were achieved. The fermented solids (biocatalyst) were employed for the selective hydrolysis of sardine oil.
Methods Fungal Strain T. lanuginosus SS-2 isolated from sugar cane bagasse (Gen Bank ID: JQ639283) was supplied by Dr. Jesus Cordova at the University of Guadalajara (Mexico). This strain was lyophilized for storage.
Appl Biochem Biotechnol
Pre-inoculum Preparation Petri dishes containing 30 mL of potato dextrose agar (PDA) covered with a cellophane membrane (Cellophane No. 5, obtained from Central Supplies in Mexico City) were inoculated with a piece of mycelium (0.5×0.5 cm) obtained from a previous culture on PDA at 45 °C for 7 days. PDA and cellophane membranes were separately sterilized at 121 °C for 15 min. Culture Media Modified Pontecorvo medium with the following composition was used for inoculum production (in g/L): (NH4)2SO4, 4.63; urea, 3.16; KH2PO4, 14; K2HPO4, 2.4; KCl, 0.8; MgSO4 · 7H2O, 0.8; yeast extract, 0.8; glucose, 28.8; olive oil, 7.2; polyvinyl alcohol, 2.1; and 1.2 mL of trace element solution containing (in mg/L) 100 Na2B4O7 · 10H2O, 50 MnCl2 · 4H2O, 50 Na2MoO4 · 2H2O, 250 CuSO4 · 5H2O, 85 FeCl3 · 6H2O, and 100 ZnSO4 · 7H2O. Culture medium for lipase production by SSF was 3-fold concentrated. Initial pH was adjusted to 6.5. Inoculum Preparation Two pieces of mycelia (0.5×0.5 cm) over cellophane membrane, obtained from a preinoculum, were cut and transferred to a sterile tube containing 5 mL of sterile culture medium and 4 g of glass bead (3 mm, Marienfeld Laboratory Glassware). Mycelium was gently disaggregated by manual agitation for 2 min. Two milliliters of the obtained suspension, free of glass bead, was added to a 125-mL Erlenmeyer flask containing 28 mL of culture medium. Flasks were incubated at 40 °C in an orbital shaker (150 rpm) for 72 h. The obtained culture was used to inoculate the culture medium for SSF. Production of Lipases by SSF Perlite was used as an inert support. It was previously washed with distilled water, dried, and sieved (mesh 18/20). Support was impregnated with inoculated (10 % v/v) culture medium (65 mL of inoculated medium and 35 g of perlite) to obtain an initial moisture content of 65 % (w/v). Glass columns for SSF (2.5-cm diameter×20 cm long) [24] were filled with 8 g of the impregnated solid medium and incubated at 45 °C for 120 h, supplying water-saturated air at a flow rate of 50 mL/min. Two columns were sampled at different time intervals for lipase, pH, and moisture content assays. CO2 production was monitored online by connecting the outlet air stream to a gas analyzer. CO2 production rate, specific CO2 production rate (μCO2), and lag phase were estimated based on the CO2 concentration in the exhaust gas stream [25]. Sample Treatment Lipase activity was assayed using enzymatic extracts or dry biocatalysts [21]. The enzymatic extracts were obtained from 1 g of dried fermented material suspended in 10 mL of distilled water, agitated in vortex for 2 min, and centrifuged at 10,000 rpm for 3 min. Supernatants were kept at 4 °C for further assays (lipase activity, pH, and glucose). The dry biocatalysts were obtained by blowing air (at 10 L/min) into a glass column containing the fermented solid to a final moisture content below 1 % w/v. The dry fermented solids were milled with a mortar to obtain a fine powder that was stored at room temperature in the dark. This dry biocatalyst maintained its full activity for at least 3 months. Moisture content was determined by using a thermobalance (Ohaus Model MB45, USA).
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Enzyme Immobilization For the hydrolysis studies of sardine oil, lipases from the dry biocatalyst were extracted and immobilized as follows: In order to get similar enzymatic activity for the preparation of the biocatalyst and the octyl-agarose-immobilized enzyme, 1 g of dry biocatalyst and 10 mL of cold (4 °C) distilled water were mixed for 2 min. Supernatants were collected by centrifugation at 8,000 rpm for 10 min. Ten milliliters of supernatant were added to 1 g of octyl-agarose and maintained under mild agitation at 4 °C until no activity was detected in the filtered supernatant (1 h). In order to evaluate the loss of activity due to the immobilization procedure, the activity of the enzyme solution without support was monitored at the same conditions. Finally, the immobilized enzyme was washed three times with 10 mL of phosphate buffer (25 mM, pH 7) and stored at 4 °C. Enzymatic Activity Assays Spectrophotometric Assay The substrate solution contained one volume of 10 mM pnitrophenyl octanoate (p-NPO) (dissolved in 2-propanol) and nine volumes of 100 mM of Tris–HCl (pH 8) and 0.25 % (w/v) of polyvinyl alcohol (PVA) [19]. The enzymatic reaction was started by adding 50 μL of enzymatic extract to 100 μL of substrate solution. Absorbance was monitored at 405 nm using a multiwell-microplate reader (Biotek Elx808, USA) at 30 °C for 5 min. The reaction rate was calculated from the slope of a standard curve of absorbance versus time, using a molar extinction coefficient of 7,859/cm M) for p-nitrophenol at the assay conditions (pH 8, 30 °C). For all assays, the hydrolysis rate of a control (buffer instead of enzymatic extract) was recorded and subtracted. One unit of enzyme activity (U) was defined as the amount of enzyme releasing 1 μmol of p-nitrophenol per minute. Titrimetric Assay The substrate emulsion was prepared as follows: 10 mL of trioctanoin and 90 mL of 10 % (w/v) gum arabic at pH 7 were emulsified with a mechanical homogenizer (Oster blender with two speeds, Model 2523-13 440-20, Mexico) for 10 min. The trioctanoin emulsion (6 mL) was mixed with 24 mL of 0.3 mM Tris–HCl (pH 7) containing 150 mM NaCl and 2 mM CaCl2. The mixture was pre-incubated at the selected temperature, and the reaction was started by adding the enzymatic extract (adequately diluted) or the biocatalyst. Initial pH was set at 7, and the released fatty acids were continuously titrated with 0.1 M NaOH for 5 min, using an automatic titrator (Mettler DL21 Titrator, Switzerland). Each assay was performed by triplicate. One unit of enzymatic activity (U) was defined as the amount of enzyme releasing 1 μmol of fatty acid per minute [19]. Possible proteases contained in the enzymatic extracts were inhibited using a cocktail specific for fungal proteases (P8215; Sigma); for that, 10 μL of the protease inhibitor (2 mM) was added to 50 μL of enzymatic extract, and the lipase activity was spectrophotometrically estimated using p-NPO at 30 °C and pH 8. Effect of Temperature on Lipase Activity and Stability The effect of temperature on lipase activity was studied using the titrimetric assay. Trioctanoin emulsions were pre-incubated at temperatures from 30 to 85 °C. To start the enzymatic reaction, 50 mg of biocatalyst was added and continuously titrated with 0.1 M NaOH for 5 min under mild agitation. Lipase activity was assayed by triplicate, and the obtained values were used to estimate activation (Ea) and deactivation (Ed) energy, according to the Arrhenius equation. The lipase stability was evaluated using the spectrophotometric assay with p-NPO as
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substrate. Enzymatic extracts were pre-incubated at temperatures from 30 to 80 °C for 4 h, taking samples every 30 min and assaying the residual lipase activity at 30 °C. Effect of pH on Lipase Activity and Stability The effect of pH on the lipase activity at 30 °C and stability at 4 °C was evaluated spectrophotometrically at 348 nm. Enzymatic extracts with 100 mM p-NPO as substrate (for activity assays) or without p-NPO (for stability assays) were incubated at different pH values. For that, the following buffers were used (at 0.1 M): citrate–phosphate, pH 3 to 6, Tris–HCl, pH 7 to 9, NaHCO3–NaOH, pH 10–11, and KCl- NaOH, pH 12. For enzymatic stability, residual activity was spectrophotometrically assayed at pH 8 and 30 °C for 5 min. Hydrolysis of Sardine Oil Sardine oil was hydrolyzed following the methodology proposed by Fernández-Lorente et al. [4]. One gram of dry biocatalyst or octyl-immobilized biocatalyst was added to a 15-mL flask containing 10 mL of a biphasic mixture of 0.1 mL of sardine oil in 4.9 mL of cyclohexane/ 5 mL of aqueous buffer solution (10-mM phosphate buffer for pH 6 or 10 mM Tris–HCl for pH 8). The flasks were covered and maintained under gentle magnetic stirring at different temperatures for different time intervals. One unit was defined as the amount of enzyme required to release 1 μmol of EPA and/or DHA per minute under the above conditions. Experiments were carried out by duplicate with coefficients of variation lower than 5 %. Analysis of EPA and DHA by HPLC-UV Samples were taken at different time intervals and used to determine the released PUFAs; for that, 50 μL of sample was dissolved in 400 μL of acetonitrile and analyzed by HPLC (Spectra Physic HPLC-UV) with an UV detector (SP 100–UV detector SP 8450) and equipped with a Kromasil C8 (15×0.4 mm) column. Samples were eluted with acetonitrile/water/acetic acid (70:30:0.1) at pH 3 as mobile phase at an isocratic flow rate of 1 mL/min. PUFAs were detected at 215 nm. Retention times were 10.3 min for EPA and 13.2 min for DHA.
Results and Discussions Kinetics of CO2 Production of T. lanuginosus Cultured on SSF Kinetics of CO2 production and CO2 production rate are shown in Fig. 1. The lag phase of 13.4 h was similar to that reported for submerged cultures of T. lanuginosus [26] and a recombinant strain of A. niger, expressing a lipase-encoding gene from T. lanuginosus [27]. Maximum CO2 production rate (3.7 mg CO2/g initial dry matter (idm) h) was reached after 31 h of culture. At the end of culture (120 h), 116 mg CO2/g idm was produced (Fig. 1a). It is worth to mention that due to the difficulties to estimate the microbial growth on solid cultures, kinetic parameters of T. lanuginosus on SSF have not been reported. The specific CO2 production rate obtained in this study (0.11/h) is slightly lower than that reported for the recombinant strain of A. niger, expressing the lipase of T. lanuginosus [27]. In related work, Gomes et al. [26] found specific growth rates from 0.1 to 0.31/h for T. lanuginosus growing in submerged cultures. On the other hand, high specific growth rate values (0.89 and 0.46/h) were observed during the biphasic growth of T. lanuginosus in rich YPS medium [28].
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Lipase Production by T. lanuginosus Cultured on SSF
120
4 a) 3
80 2 40 1
0
Production of CO2 (mg/g idm)
CO2 production rate (mg/g idm h)
Although glucose is a well-recognized catabolic repressor, previous studies report low catabolite repression during the production of pectinases [29] and tannases [30] by A. niger cultured on SSF. In this study, lipase production started once glucose was completely consumed after 24 h of culture (results not shown), reaching a maximum value of 22±2.2 U/g dm at 48 h of culture (Fig. 1b) and remaining nearly constant (19.5±1.8 U/g dm) until the end of culture (120 h). The use of the Luedecking–Piret equation [31] allowed showing that lipase production was directly associated to CO2 production (α=0.244 U/mg) with a nonassociated level (β=−9× 10−4 U/mg h) nearly negligible at the end of culture (Table 1). The high absolute value of η (−30) suggested a high degree of correlation between lipase production and CO2 production [18].
0
0
24
48
72
96
120
Time (h)
Lipase Activity (U/g dm)
30
b)
25 20 15 10 5 0 0
24
48
72 Time (h)
96
120
Fig. 1 Kinetics of CO2 production (mg/g idm—points) and CO2 production rate (mg/g idm h—continuous line) (a) and lipase production (b) of T. lanuginosus cultured on SSF at 40 °C, using perlite as inert support impregnated with a culture medium
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Table 1 Kinetic parameters of T. lanuginosus growth cultured on SSF
α production of enzymatic activity associated to the production of CO2, η ratio αμ/β, β rate of enzymatic activity not associated to the production of CO2
Parameter
Value
CO2max (mg/g idm)
116.4±10.7
CO2 (mg/g idm h) μ (1/h)
3.5±0.4 0.11±0.01
Lag phase (h)
13.4±0.5
α (U/mg CO2)
0.244
β (U/mg CO2 h)
−0.0009
η
−30
Additionally, the low β value suggested the absence of proteases at the end of culture. This was corroborated assaying lipase activity in the presence of a proteases inhibitor; finding that activity remained constant with or without the inhibitor. Catalytic characteristics of the biocatalysts produced in this work were carried out using either enzymatic extracts or dry biocatalysts. For that, SSF samples were obtained at the same physiological state, namely, once the maximum CO2 production rate was achieved (around 44 h of culture, see Fig. 2a). The physiological state of culture was monitored by online analysis of CO2 in the exhaust air phase. Lipase activity was titrimetrically assayed with trioctanoin for both the enzymatic extract (liquid) and the biocatalyst (solid) samples, obtaining 58±1 and 63±2 U/g dm, respectively. Lipase activity was also assayed with the spectrophotometric method with p-nitrophenyl octanoate as substrate. Twenty times higher activity was obtained with the titrimetric method. Likewise, measurement of lipase activity with olive oil as substrate yielded enzymatic titers up to 13.5 higher than that obtained with p-nitrophenyl palmitate as substrate [24]. Effect of Temperature on Lipase Activity The effect of temperature on lipase activity of T. lanuginosus was evaluated by incubating the enzymatic extracts from 30 to 85 °C. Lipase activity increased up to 2.0-fold from 30 to 60 °C and at higher incubation temperatures (from 60 to 85 °C); the lipase activity remained nearly constant (Fig. 2a). To our knowledge, 85 °C has been the highest record reported for lipases produced by filamentous fungi (Table 2). Similar profile of lipase activity as a function of temperature was obtained using the solid biocatalyst and the titrimetric assay (results not shown). Lipase activity at 85 °C is higher than the reported (80 °C) for the well-known immobilized lipase B of Candida antarctica [32]. High temperatures have also been reported (70 °C) for xylanases and (70 °C) for phytases from T. lanuginosus [33]. The calculated activation energy (Ea=27 kJ/mol) for the lipase produced in this work was similar to those reported for the lipases from Candida rugosa (30 to 40 kJ/mol) [34] and from Rhizopus homothallicus (30 kJ/mol) [19] (Table 3). However, higher Ea value was reported (124.8 kJ/mol) for xylanases from T. lanuginosus [36]. The calculated deactivation energy (Ed=6.7 kJ/mol) for the lipase produced in this work was lower than those reported for a commercial lipase from T. lanuginosus (26.5 kJ/mol) [37] and for xylanases produced by wild and mutants strains of T. lanuginosus (98.6 and 71.8 kJ/mol, respectively) [36]. The high Ea/ Ed ratio obtained herein (4.0) gives evidence about the potential application of the biocatalyst at high temperatures of reaction [38, 39]; for instance, the acidolysis reactions for the production of structured lipids are carried out from 45 to 50 °C [40].
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Lipase activity (U/g dm)
150
a)
120 90 60 30 0 30
Relative activity (%)
120
40
50 60 70 Temperature ( C)
80
90
b)
90
60
30
0 3
4
5
6
7
8
9
10
11
12
pH Fig. 2 Effect of temperature (a) and pH (b) on lipase activity of T. lanuginosus produced on SSF. Error bars represent the standard deviation of three independent assays. Activity was titrimetrically assayed with trioctanoin as substrate
Effect of Temperature on Lipase Stability The thermostability studies carried out with the enzymatic extracts of T. lanuginosus showed that lipase activity was enhanced from 30 to 50 °C and maintained constant at 60 °C, after an incubation time of 4 h. At 70 °C, enzyme activity decreased 50 % after being incubated in 0.5 h. An increase of activity up to 30 % was observed from 30 to 60 °C after 2 h of incubation. Some authors have discussed the high thermal stability and improvement in
Appl Biochem Biotechnol Table 2 Effect of temperature on activity and stability of lipases produced by solid-state fermentation (SSF) and submerged fermentation (SmF) for different fungi strains Fungi
Topt (°C)
T* (°C)
T. lanuginosus
80
70
0.5
80
80
0.25
30
50
0.4
nr
SmF
40
50
0.7
nr
SSF
37
50
0.7
1.0
cp
50
50
1.1
0.6
cp
Penicillium sp.
37
65
1.5
1.9
SSF
Humicola lanuginosa
42 45
25 70
0.3 ≈2
