Enzyme and Microbial Technology 71 (2015) 8–12
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Enzyme and Microbial Technology journal homepage: www.elsevier.com/locate/emt
A convenient test for lipase activity in aqueous-based solutions Jin Guo a , Cheng-Peng Chen b , Shu-Gen Wang a,∗ , Xiao-Jun Huang b,∗∗ a
Key Laboratory of Eco-Textile (Jiangnan University), Ministry of Education, Wuxi, Jiangsu 214122, China MOE Key Laboratory of Macromolecular Synthesis and Functionalization, Department of Polymer Science and Engineering, Zhejiang University, Hangzhou 310027, China b
a r t i c l e
i n f o
Article history: Received 29 August 2014 Received in revised form 12 January 2015 Accepted 19 January 2015 Available online 25 January 2015 Keywords: Candida rugosa lipase p-nitrophenyl palmitate Lipase enzymatic activity measurement Uniform aqueous-based solution
a b s t r a c t We proposed a convenient and accurate method for the measurement of lipase activity in a uniform aqueous-based substrate solution. In this work, lipase from Candida rugosa was used as the model lipase to test its catalytic ability toward p-nitrophenyl palmitate (p-NPP), which was suspended in a mixture of p-NPP ethanol solution and buffer. An ultraviolet–visible spectrophotometer was used to efficiently measure the liberated p-nitrophenol without extraction or centrifugation. Several factors that affected lipase activity were investigated, such as the ratio of p-NPP ethanol solution to buffer, the concentrations of p-NPP and lipase, as well as the temperature, reaction time, pH and agitation rate. Additionally, enzyme catalytic parameters such as Km , Vm and “activation energy” were also assessed. We determined the optimal conditions for lipase in this homogeneous system and demonstrated lipase’s catalytic performance in this condition followed Michealis–Menten kinetics. © 2015 Published by Elsevier Inc.
1. Introduction Lipases, or triacylglycerol acyl ester hydrolases (EC 3.1.1.3), are enzymes possessing an intrinsic capacity to catalyze the cleavage of carboxyl ester bonds in tri-, di-, and monoacylglycerols (the major constituents of animal, plant, and microbial fats and oils) [1]. Generally, lipase can catalyze a wide range of reactions, such as hydrolysis, transesterification, alcoholysis, acidolysis and esterification [2–5]. More than 50 lipases have been identified and they have tremendous application potential in food industry, medicine, hygiene, chemistry, chemical engineering, environmental protection and energy development [6–10]. For lipases practical applications in industrial, their catalytic activity must be understood. Thus, establishing convenient and accurate methods for measuring lipase activity is of vital importance. The catalysis systems of lipase may be divided into two categories: aqueous-based mediums and organic mediums [11–13]. Commonly, lipase is a water-soluble enzyme, with good solubility
∗ Corresponding author at: Key Laboratory of Eco-Textile (Jiangnan University), Ministry of Education, 1800 Li Hu Road, Wuxi, Jiangsu 214122, China. Tel.: +86 13812089771; fax: +86 051013812089771. ∗∗ Corresponding author at: Key Laboratory of Macromolecular Synthesis and Functionalization, Department of Polymer Science and Engineering, Zhejiang University, 38 Zhe Da Road, Hangzhou 310027, China. Tel.: +86 571 87952605; fax: +86 571 87951773. E-mail addresses: [email protected] (S.-G. Wang), [email protected] (X.-J. Huang). http://dx.doi.org/10.1016/j.enzmictec.2015.01.005 0141-0229/© 2015 Published by Elsevier Inc.
and stability in the aqueous phase. However, the substrates of lipase are often insoluble in water. Thus, substrates should first be dissolved in organic solvents and subsequently mixed with buffer. According to previous reports, the substrate mixtures prepared by this method are always two-phase systems, and additives are included in order to improve the uniformity and stability of these metastable mixtures [14,15]. However, many studies have reported that surfactants influence lipase activity, with various activation or inhibition effects [16–18]. In organic mediums, lipases are usually directly added to an organic solvent that also contains dissolved substrate [19]. This method is advantageous because the substrate is soluble and the mixture is stable. Unfortunately, protein molecules have both denature feature and conformational flexibility, which lead to their activity reduce in the presence of organic solvents [20,21]. Based on the discussion above, it is necessary to design a suitable method for the measurement of lipase activity. Our study focused on the design of an aqueous-based substrate solution without additives for the determination of lipase catalytic properties. We test several solvents and ethanol was chosen as an appropriate solvent to dissolve p-NPP for the following reasons: first, it is miscible with water at any volume fraction. Second, p-NPP dissolves well in ethanol [14]. Finally, ethanol is a relative high polarity organic solvent and has less influence on lipase conformation compared with 2-propanol, which is a solvent of the substrate in previously reported methods [22]. In this work, a uniform solution was prepared by mixing a p-NPP ethanol solution with a certain volume of buffer at 65 ◦ C. Then, this solution was equilibrated at a certain temperature and subsequently used for
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the measurement of enzymatic activity. Moreover, lipase from Candida rugosa was used to investigate the hydrolysis of p-NPP in this particular aqueous-based solution. After the enzyme-catalytic reaction, the amount of liberated product in the uniform solution was directly measured with an ultraviolet–visible (UV) spectrophotometer. Factors that affect enzyme activity, such as the volume proportion of water/ethanol, reaction time, p-NPP and lipase concentrations, temperature, pH and agitation rate were all investigated in this work. In addition, we also determined the catalytic parameters in this aqueous-based solution. 2. Materials and methods 2.1. Materials Lipase powder from C. rugosa (1150 units/mg solid), Bradford reagent, bovine serum albumin (BSA, molecular mass: 67,000 Da) and p-NPP were obtained from Sigma–Aldrich Chemical Co. (St. Louis, MO, USA) and used as received. KH2 PO4 , Na2 HPO4 , NaCl and KCl were obtained from Chemsynlab Pharmaceutical Science & Technology (Beijing, China). All other chemicals were of analytical grade and used without further purification. The water used in all experiments was deionized and ultrafiltered in order to obtain a resistance of 18 M cm with a TKA MicroPure water system. The phosphate buffer solution (PBS, 0.05 M) was prepared with NaCl (8.5 mg/mL), Na2 HPO4 (2.2 mg/mL) and NaH2 PO4 (0.4 mg/mL), and NaOH/HCl was used to adjust the pH. 2.2. Preparation of the homogeneous aqueous-based solution Following standard protocols, p-NPP was dissolved in ethanol in a 50-mL Erlenmeyer flask and heated to 65 ◦ C to form the p-NPP solution at the concentrations of 0.2, 0.4, 0.6, 0.8, 1.0, 1.5, 2.0 mg/mL. Then, PBS (0.05 M) of 65 ◦ C was cautiously and dropwise added. The resulting mixture was in a metastable state and the soluble pNPP molecular may aggregate with the storage time and the change of temperature. The prepared mixture should be used timely in order to avoid the phase separation. Dynamic light scattering (DLS) was used to characterize the substrate states in this media stored at room temperature at several different equilibration times. 2.3. Analysis of lipase secondary structure via Circular Dichroism (CD) spectroscopy Circular Dichroism (CD) spectroscopy was used for the examination of lipase structures in different solutions, which were recorded on a spectropolarimeter (MOS-450 AF/AF-CD, Bio-Logic, France) using a 0.2 cm quartz cuvette. For those three lipase different solution, lipase powder (3 mg/mL) was diluted in PBS (pH 7.0, 0.05 M), ethanol/PBS (pH 7.0, 0.05 M) (v:v, 1:1) and 2-propanol/PBS (pH 7.0, 0.05 M) (v:v, 1:1), respectively. Then, those solutions were centrifuged at 400 rpm for 5 min to remove insoluble impurities. The cell holder compartment was maintained in a nitrogen atmosphere at room temperature. At least five spectra were measured and the average was recorded. 2.4. Lipase activity measurement with a UV-spectrophotometer An aqueous enzyme solution (PBS, 0.05 M, pH 7.0) of lipase (3 mg/mL) was centrifuged at 400 rpm for 5 min to remove any insoluble impurities. The protein concentration in the solution was determined using Coomassie Brilliant Blue reagent, following Bradford’s method [23]. BSA was used as a standard to construct a calibration curve. The prepared homogeneous solution was pre-equilibrated at a determined temperature for a certain time. The catalytic reaction was initiated by adding an enzyme solution. Sodium carbonate solution (0.5 M) of the same volume was added to terminate the reaction (the pH value for this solution was 11.8) and enhance the detection sensitivity after the catalytic reacted for a certain time. Finally, the lipase catalytic activity was calculated from the absorbance of this mixture at an appropriate dilution in phosphate buffer against the blank without enzyme using a UV spectrophotometer (UV-1610, Shimadzu. Japan) at 410 nm. This was measured as one enzyme unit was the amount of protein liberating 1.0 mol p-NP/min under these conditions. Each treatment was measured in at least three parallel experiments and the average was recorded.
3. Results and discussion 3.1. Solubility of p-NPP in ethanol In this study the hydrolysis of p-NPP by C. rugosa lipase was used as a test reaction. p-NPP was dissolved in ethanol and the solubility of p-NPP was investigated. As shown in Fig. 1, the solubility of p-NPP in ethanol conformed to Eq. (1). “S” indicates the highest weight (g)
Fig. 1. Solubility of p-NPP in ethanol, measured by change temperature.
of p-NPP dissolved in 100 g ethanol, which increased as the temperature increased. The intersection of the tangent of the curve indicated that the solubility mutation temperature was nearly 41 ◦ C. At room temperature p-NPP solubility in 100 g ethanol solvent was approximately 0.52 g (2.44 mg/mL) at room temperature. S = 0.08985 exp
T
11.61
− 0.2584
(1)
We found that ethanol was a satisfactory organic solvent for lipase. However, organic molecules may have an effect on lipase secondary structures, CD spectroscopy was used to examine changes of lipase secondary structure in different organic solvent/buffer (v:v = 1:1) mixtures (Fig. S1). The molecules spectra in the far ultraviolet regions were dominated by the n → p* and p → p* transitions of amide groups, which was influenced by the geometries of the polypeptide backbones, their spectra were reflective of the different types of secondary structures present [24]. Obviously, the secondary structures of lipase in ethanol/PBS and 2-propanol/PBS mixture differed from the structure in PBS. Lipase underwent less structural change in ethanol/PBS than in the isopropanol/PBS mixture. DLS measurements revealed that this substrate remains clear and uniform for 1 h, which was enough to measure lipase activity. We concluded that ethanol is a suitable organic solvent for the measurement of lipase activity in aqueousbased solutions. 3.2. Effect of PBS volume on lipase reaction rate The purpose of this research was to design a uniform aqueous solution without additives that would be optimal for the measurement of lipase activity. As described above, we dissolved a certain amount of substrate (p-NPP) in ethanol and then mixed it with PBS. The optimum ratio of p-NPP ethanol solution and PBS was dependent on many factors, including solvent and substrate properties and reaction temperature. We also investigated the catalytic properties of lipase in different volume percentages of PBS (0.05 M, pH 7.0). A bell-shaped curve of enzyme activity was obtained as PBS volume increased (Fig. S2). Lipase catalytic activity firstly increased until the peak point at 50% with the maximal reaction rate of 1.337 ± 0.081 U mg−1 . Then, it reduced in high water content because of the substrate state transformed to small solid particles by aggregation, which might cause p-NPP diffusion limitation [25]. It was obvious that lipase showed low activity in neat ethanol, which was consistent with previous reports [21]. In most cases, lipase enzyme catalytic efficiency in
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Fig. 2. Effect of pH on the activity of lipase.
neat organic solvent was several orders of magnitude lower than in aqueous systems. This behavior could be ascribed to different causes, such as limited diffusion limitations, high saturating substrate concentrations and restricted protein flexibility. 3.3. Effect of pH on lipase performance The pH value of the aqueous-based solution was an important parameter that affected the rate and conversion of lipase-catalyzed activity. The hydrolysis of p-NPP was assayed by varying buffer pH from 5.0 to 9.0. We observed that the optimum pH for lipase was about 5.5 (Fig. 2), close to the isoelectric point (pI) of C. rugosa lipase. The highest hydrolysis conversion rates occurred in these conditions with a lipase activity of 4.023 ± 0.372 U mg−1 . 3.4. Effect of reaction temperature on lipase activity Fig. 3(A) shows the effect of temperature on the hydrolysis rate of p-NPP by lipase in this homogeneous aqueous-based solution. It is conceivable that the lipase activity increased when the temperature increased from 20 to 45 ◦ C, and lipase’s optimum temperature of this system was approximately 45 ◦ C with the activity of 4.02 ± 0.37 U mg−1 . It should be noted that a certain amount of “activation energy” (Ea ) was necessary to start the chemical reactions. Enzymatic reactions may be optimized to accelerate the reaction rate by reducing the activation energy. It was found that the activation energy of enzyme catalytic system related to many factors, such as the lipase species of origin, the amount of lipase added to the solution, the state of the substrate as well as the solvent properties. In order to investigate the Ea value of lipase in a homogeneous aqueous-based solution, we re-assessed Fig. 3(A). Results shown that the plots in Fig. 3(B) followed a classic Arrhenius-law function and had an Ea value of 31.7 kJ/mol. This result is generally within the scope that such reactions have activation energies in the range of 20–40 kJ/mol [26–29].
Fig. 3. (A) Effect of temperature on performance of lipase. (B) Arrhenius plot obtained from the first part of the reaction rate versus temperature (20–45 ◦ C).
Fig. 4. (A) Effect of the p-NPP concentration on its hydrolysis reaction rate catalyzed by lipase of C. rugosa in ethanol/buffer solution. (B) Lineweaver–Burk plot.
3.5. Determination of Km and Vm values Kinetics of the hydrolytic reaction was investigated using initial p-NPP concentration range of 0–2.0 mg/mL (0–2.16 mM) in the homogeneous solutions. As shown in Fig. 4(A), the hydrolytic rate did not significantly increase after the p-NPP concentration reached 1.0 mg/mL, which was about 1.29 ± 0.05 U mg−1 . Fig. 4(C) shows
the reaction mixture quantity as a function of substrate concentration. The amount of the product increased with the increase of the p-NPP’s concentration, and the color of the solution was also gradually deepened. The kinetic parameters as Km and Vm values were studied, and the result is shown in Fig. 4(B). Lineweaver–Burk plots were used to calculate the kinetics parameters. Km and Vm values of
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Table 1 Kinetic parameters (Vm and Km ) of lipase in an aqueous-based solution. Lipase origin
Reaction mixture
Vm (U mg−1 )
Km (mM)
Reference
Candida rugosa Candida rugosa Rhizomucor miehei Pseudomonas cepacia
Ethanol/PBS Ethanol/PBS n-Hexane n-Heptane
1.99 3.98 11.72 30
0.76 13 3.03 12
This work [28] [29] [19]
(correlation coefficient (R2 ) of 0.97) with different amount of lipase added. This linear relationship decreased with time because the substrate was gradually consumed. 4. Conclusion
Fig. 5. Kinetics of p-NPP hydrolysis with different amounts (3.56 × 10−3 , 5.34 × 10−3 , 7.13 × 10−3 , 10.69 × 10−3 , 14.25 × 10−3 mg/mL) of lipase and different reactions times (3, 5, 10, 15 and 25 min) under the optimal reaction conditions.
This research centered on design a uniform aqueous-based solution appropriate for the measurement of lipase’s activity. Additionally, the effect of the solution properties on the catalytic activity of C. rugosa lipase was also investigated. To prepare a homogeneous reaction mixture, p-NPP was first dissolved in ethanol at a concentration of 1 mg/mL, and then mixed with PBS at a volume ratio of 1:1. In this aqueous-based solution, the reaction rate was found to be maximum at 45 ◦ C and the pH value of 5.5. In this aqueous solution lipase follows Michealis–Menten saturation kinetics, Vm and Km values are 1.99 U mg−1 and 0.76 mM, respectively. The activation energy of lipase in this solution was 31.71 kJ/mol. This measurement method of lipase catalytic activity in this aqueousbased solution was more convenient and accurate in comparison with previously reported methods. Acknowledgments
different types of lipases in different reaction systems are also listed and compared in Table 1. It was clear that the Km value of lipase in this system (0.76 mM) was lower than Km value in emulsified systems (13 mM) [29]. Vm (maximum reaction rate), which defines the highest possible velocity when all the enzyme is saturated with substrate, reflects the intrinsic characteristics of the enzyme, but may be affected by diffusion constrains [19,30]. The value of this system was also got from the inserted figure in Fig. 4 and the result was 1.99 U/mg.
The authors would like to thank the financial support from the National Natural Science Foundation of China (Grant No. 21274126 and No. 51473143), National “Twelfth Five-Year” Plan for Science & Technology Support of China (No. 2012BAI08B01). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.enzmictec.2015. 01.005.
3.6. Optimal conditions for lipase activity assays On the basis of the experimental results described above, the optimum conditions for lipase catalytic reactions are listed as follows: a buffer/ethanol volume ratio of 1:1, reaction temperature of 45 ◦ C, pH value of 5.5 and p-NPP content of ethanol solution of 1 mg/mL. The process to prepare the reaction system followed the description above. The volume of PBS and p-NPP’s ethanol solution was 1 mL. The amount of p-NPP hydrolysis was measured under conditions with different amounts of enzyme added (7.125 × 10−3 , 10.6875 × 10−3 , 14.25 × 10−3 , 21.375 × 10−3 , 28.5 × 10−3 mg) and different catalytic times (3, 5, 10, 15 and 25 min). As shown in Fig. 5(A), the absorbance was gradually increased with p-NPP hydrolyzed and the substrate was almost completely hydrolyzed after 30 min. It was obvious the more amount of enzyme added the faster the initial hydrolysis rate, and the shorter time required to completely hydrolysis the substrate. Moreover, as the initial content of p-NPP in this reaction solution was 1 mg/mL (2.65 mM) and the liberated p-NP concentration is 2.25 mM after hydrolyzed of 5 min at the lipase concentration of 14.25 × 10−3 mg/mL, so it was nearly 85% of the substrate being hydrolyzed. Fig. 5(B) shows the reaction rates of different lipase addition in different times (3, 5 and 10 min). The initial reaction rates of lipase added amounts of the first 3 min increased nearly linearly
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Contents lists available at ScienceDirect
Enzyme and Microbial Technology journal homepage: www.elsevier.com/locate/emt
A convenient test for lipase activity in aqueous-based solutions Jin Guo a , Cheng-Peng Chen b , Shu-Gen Wang a,∗ , Xiao-Jun Huang b,∗∗ a
Key Laboratory of Eco-Textile (Jiangnan University), Ministry of Education, Wuxi, Jiangsu 214122, China MOE Key Laboratory of Macromolecular Synthesis and Functionalization, Department of Polymer Science and Engineering, Zhejiang University, Hangzhou 310027, China b
a r t i c l e
i n f o
Article history: Received 29 August 2014 Received in revised form 12 January 2015 Accepted 19 January 2015 Available online 25 January 2015 Keywords: Candida rugosa lipase p-nitrophenyl palmitate Lipase enzymatic activity measurement Uniform aqueous-based solution
a b s t r a c t We proposed a convenient and accurate method for the measurement of lipase activity in a uniform aqueous-based substrate solution. In this work, lipase from Candida rugosa was used as the model lipase to test its catalytic ability toward p-nitrophenyl palmitate (p-NPP), which was suspended in a mixture of p-NPP ethanol solution and buffer. An ultraviolet–visible spectrophotometer was used to efficiently measure the liberated p-nitrophenol without extraction or centrifugation. Several factors that affected lipase activity were investigated, such as the ratio of p-NPP ethanol solution to buffer, the concentrations of p-NPP and lipase, as well as the temperature, reaction time, pH and agitation rate. Additionally, enzyme catalytic parameters such as Km , Vm and “activation energy” were also assessed. We determined the optimal conditions for lipase in this homogeneous system and demonstrated lipase’s catalytic performance in this condition followed Michealis–Menten kinetics. © 2015 Published by Elsevier Inc.
1. Introduction Lipases, or triacylglycerol acyl ester hydrolases (EC 3.1.1.3), are enzymes possessing an intrinsic capacity to catalyze the cleavage of carboxyl ester bonds in tri-, di-, and monoacylglycerols (the major constituents of animal, plant, and microbial fats and oils) [1]. Generally, lipase can catalyze a wide range of reactions, such as hydrolysis, transesterification, alcoholysis, acidolysis and esterification [2–5]. More than 50 lipases have been identified and they have tremendous application potential in food industry, medicine, hygiene, chemistry, chemical engineering, environmental protection and energy development [6–10]. For lipases practical applications in industrial, their catalytic activity must be understood. Thus, establishing convenient and accurate methods for measuring lipase activity is of vital importance. The catalysis systems of lipase may be divided into two categories: aqueous-based mediums and organic mediums [11–13]. Commonly, lipase is a water-soluble enzyme, with good solubility
∗ Corresponding author at: Key Laboratory of Eco-Textile (Jiangnan University), Ministry of Education, 1800 Li Hu Road, Wuxi, Jiangsu 214122, China. Tel.: +86 13812089771; fax: +86 051013812089771. ∗∗ Corresponding author at: Key Laboratory of Macromolecular Synthesis and Functionalization, Department of Polymer Science and Engineering, Zhejiang University, 38 Zhe Da Road, Hangzhou 310027, China. Tel.: +86 571 87952605; fax: +86 571 87951773. E-mail addresses: [email protected] (S.-G. Wang), [email protected] (X.-J. Huang). http://dx.doi.org/10.1016/j.enzmictec.2015.01.005 0141-0229/© 2015 Published by Elsevier Inc.
and stability in the aqueous phase. However, the substrates of lipase are often insoluble in water. Thus, substrates should first be dissolved in organic solvents and subsequently mixed with buffer. According to previous reports, the substrate mixtures prepared by this method are always two-phase systems, and additives are included in order to improve the uniformity and stability of these metastable mixtures [14,15]. However, many studies have reported that surfactants influence lipase activity, with various activation or inhibition effects [16–18]. In organic mediums, lipases are usually directly added to an organic solvent that also contains dissolved substrate [19]. This method is advantageous because the substrate is soluble and the mixture is stable. Unfortunately, protein molecules have both denature feature and conformational flexibility, which lead to their activity reduce in the presence of organic solvents [20,21]. Based on the discussion above, it is necessary to design a suitable method for the measurement of lipase activity. Our study focused on the design of an aqueous-based substrate solution without additives for the determination of lipase catalytic properties. We test several solvents and ethanol was chosen as an appropriate solvent to dissolve p-NPP for the following reasons: first, it is miscible with water at any volume fraction. Second, p-NPP dissolves well in ethanol [14]. Finally, ethanol is a relative high polarity organic solvent and has less influence on lipase conformation compared with 2-propanol, which is a solvent of the substrate in previously reported methods [22]. In this work, a uniform solution was prepared by mixing a p-NPP ethanol solution with a certain volume of buffer at 65 ◦ C. Then, this solution was equilibrated at a certain temperature and subsequently used for
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the measurement of enzymatic activity. Moreover, lipase from Candida rugosa was used to investigate the hydrolysis of p-NPP in this particular aqueous-based solution. After the enzyme-catalytic reaction, the amount of liberated product in the uniform solution was directly measured with an ultraviolet–visible (UV) spectrophotometer. Factors that affect enzyme activity, such as the volume proportion of water/ethanol, reaction time, p-NPP and lipase concentrations, temperature, pH and agitation rate were all investigated in this work. In addition, we also determined the catalytic parameters in this aqueous-based solution. 2. Materials and methods 2.1. Materials Lipase powder from C. rugosa (1150 units/mg solid), Bradford reagent, bovine serum albumin (BSA, molecular mass: 67,000 Da) and p-NPP were obtained from Sigma–Aldrich Chemical Co. (St. Louis, MO, USA) and used as received. KH2 PO4 , Na2 HPO4 , NaCl and KCl were obtained from Chemsynlab Pharmaceutical Science & Technology (Beijing, China). All other chemicals were of analytical grade and used without further purification. The water used in all experiments was deionized and ultrafiltered in order to obtain a resistance of 18 M cm with a TKA MicroPure water system. The phosphate buffer solution (PBS, 0.05 M) was prepared with NaCl (8.5 mg/mL), Na2 HPO4 (2.2 mg/mL) and NaH2 PO4 (0.4 mg/mL), and NaOH/HCl was used to adjust the pH. 2.2. Preparation of the homogeneous aqueous-based solution Following standard protocols, p-NPP was dissolved in ethanol in a 50-mL Erlenmeyer flask and heated to 65 ◦ C to form the p-NPP solution at the concentrations of 0.2, 0.4, 0.6, 0.8, 1.0, 1.5, 2.0 mg/mL. Then, PBS (0.05 M) of 65 ◦ C was cautiously and dropwise added. The resulting mixture was in a metastable state and the soluble pNPP molecular may aggregate with the storage time and the change of temperature. The prepared mixture should be used timely in order to avoid the phase separation. Dynamic light scattering (DLS) was used to characterize the substrate states in this media stored at room temperature at several different equilibration times. 2.3. Analysis of lipase secondary structure via Circular Dichroism (CD) spectroscopy Circular Dichroism (CD) spectroscopy was used for the examination of lipase structures in different solutions, which were recorded on a spectropolarimeter (MOS-450 AF/AF-CD, Bio-Logic, France) using a 0.2 cm quartz cuvette. For those three lipase different solution, lipase powder (3 mg/mL) was diluted in PBS (pH 7.0, 0.05 M), ethanol/PBS (pH 7.0, 0.05 M) (v:v, 1:1) and 2-propanol/PBS (pH 7.0, 0.05 M) (v:v, 1:1), respectively. Then, those solutions were centrifuged at 400 rpm for 5 min to remove insoluble impurities. The cell holder compartment was maintained in a nitrogen atmosphere at room temperature. At least five spectra were measured and the average was recorded. 2.4. Lipase activity measurement with a UV-spectrophotometer An aqueous enzyme solution (PBS, 0.05 M, pH 7.0) of lipase (3 mg/mL) was centrifuged at 400 rpm for 5 min to remove any insoluble impurities. The protein concentration in the solution was determined using Coomassie Brilliant Blue reagent, following Bradford’s method [23]. BSA was used as a standard to construct a calibration curve. The prepared homogeneous solution was pre-equilibrated at a determined temperature for a certain time. The catalytic reaction was initiated by adding an enzyme solution. Sodium carbonate solution (0.5 M) of the same volume was added to terminate the reaction (the pH value for this solution was 11.8) and enhance the detection sensitivity after the catalytic reacted for a certain time. Finally, the lipase catalytic activity was calculated from the absorbance of this mixture at an appropriate dilution in phosphate buffer against the blank without enzyme using a UV spectrophotometer (UV-1610, Shimadzu. Japan) at 410 nm. This was measured as one enzyme unit was the amount of protein liberating 1.0 mol p-NP/min under these conditions. Each treatment was measured in at least three parallel experiments and the average was recorded.
3. Results and discussion 3.1. Solubility of p-NPP in ethanol In this study the hydrolysis of p-NPP by C. rugosa lipase was used as a test reaction. p-NPP was dissolved in ethanol and the solubility of p-NPP was investigated. As shown in Fig. 1, the solubility of p-NPP in ethanol conformed to Eq. (1). “S” indicates the highest weight (g)
Fig. 1. Solubility of p-NPP in ethanol, measured by change temperature.
of p-NPP dissolved in 100 g ethanol, which increased as the temperature increased. The intersection of the tangent of the curve indicated that the solubility mutation temperature was nearly 41 ◦ C. At room temperature p-NPP solubility in 100 g ethanol solvent was approximately 0.52 g (2.44 mg/mL) at room temperature. S = 0.08985 exp
T
11.61
− 0.2584
(1)
We found that ethanol was a satisfactory organic solvent for lipase. However, organic molecules may have an effect on lipase secondary structures, CD spectroscopy was used to examine changes of lipase secondary structure in different organic solvent/buffer (v:v = 1:1) mixtures (Fig. S1). The molecules spectra in the far ultraviolet regions were dominated by the n → p* and p → p* transitions of amide groups, which was influenced by the geometries of the polypeptide backbones, their spectra were reflective of the different types of secondary structures present [24]. Obviously, the secondary structures of lipase in ethanol/PBS and 2-propanol/PBS mixture differed from the structure in PBS. Lipase underwent less structural change in ethanol/PBS than in the isopropanol/PBS mixture. DLS measurements revealed that this substrate remains clear and uniform for 1 h, which was enough to measure lipase activity. We concluded that ethanol is a suitable organic solvent for the measurement of lipase activity in aqueousbased solutions. 3.2. Effect of PBS volume on lipase reaction rate The purpose of this research was to design a uniform aqueous solution without additives that would be optimal for the measurement of lipase activity. As described above, we dissolved a certain amount of substrate (p-NPP) in ethanol and then mixed it with PBS. The optimum ratio of p-NPP ethanol solution and PBS was dependent on many factors, including solvent and substrate properties and reaction temperature. We also investigated the catalytic properties of lipase in different volume percentages of PBS (0.05 M, pH 7.0). A bell-shaped curve of enzyme activity was obtained as PBS volume increased (Fig. S2). Lipase catalytic activity firstly increased until the peak point at 50% with the maximal reaction rate of 1.337 ± 0.081 U mg−1 . Then, it reduced in high water content because of the substrate state transformed to small solid particles by aggregation, which might cause p-NPP diffusion limitation [25]. It was obvious that lipase showed low activity in neat ethanol, which was consistent with previous reports [21]. In most cases, lipase enzyme catalytic efficiency in
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Fig. 2. Effect of pH on the activity of lipase.
neat organic solvent was several orders of magnitude lower than in aqueous systems. This behavior could be ascribed to different causes, such as limited diffusion limitations, high saturating substrate concentrations and restricted protein flexibility. 3.3. Effect of pH on lipase performance The pH value of the aqueous-based solution was an important parameter that affected the rate and conversion of lipase-catalyzed activity. The hydrolysis of p-NPP was assayed by varying buffer pH from 5.0 to 9.0. We observed that the optimum pH for lipase was about 5.5 (Fig. 2), close to the isoelectric point (pI) of C. rugosa lipase. The highest hydrolysis conversion rates occurred in these conditions with a lipase activity of 4.023 ± 0.372 U mg−1 . 3.4. Effect of reaction temperature on lipase activity Fig. 3(A) shows the effect of temperature on the hydrolysis rate of p-NPP by lipase in this homogeneous aqueous-based solution. It is conceivable that the lipase activity increased when the temperature increased from 20 to 45 ◦ C, and lipase’s optimum temperature of this system was approximately 45 ◦ C with the activity of 4.02 ± 0.37 U mg−1 . It should be noted that a certain amount of “activation energy” (Ea ) was necessary to start the chemical reactions. Enzymatic reactions may be optimized to accelerate the reaction rate by reducing the activation energy. It was found that the activation energy of enzyme catalytic system related to many factors, such as the lipase species of origin, the amount of lipase added to the solution, the state of the substrate as well as the solvent properties. In order to investigate the Ea value of lipase in a homogeneous aqueous-based solution, we re-assessed Fig. 3(A). Results shown that the plots in Fig. 3(B) followed a classic Arrhenius-law function and had an Ea value of 31.7 kJ/mol. This result is generally within the scope that such reactions have activation energies in the range of 20–40 kJ/mol [26–29].
Fig. 3. (A) Effect of temperature on performance of lipase. (B) Arrhenius plot obtained from the first part of the reaction rate versus temperature (20–45 ◦ C).
Fig. 4. (A) Effect of the p-NPP concentration on its hydrolysis reaction rate catalyzed by lipase of C. rugosa in ethanol/buffer solution. (B) Lineweaver–Burk plot.
3.5. Determination of Km and Vm values Kinetics of the hydrolytic reaction was investigated using initial p-NPP concentration range of 0–2.0 mg/mL (0–2.16 mM) in the homogeneous solutions. As shown in Fig. 4(A), the hydrolytic rate did not significantly increase after the p-NPP concentration reached 1.0 mg/mL, which was about 1.29 ± 0.05 U mg−1 . Fig. 4(C) shows
the reaction mixture quantity as a function of substrate concentration. The amount of the product increased with the increase of the p-NPP’s concentration, and the color of the solution was also gradually deepened. The kinetic parameters as Km and Vm values were studied, and the result is shown in Fig. 4(B). Lineweaver–Burk plots were used to calculate the kinetics parameters. Km and Vm values of
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Table 1 Kinetic parameters (Vm and Km ) of lipase in an aqueous-based solution. Lipase origin
Reaction mixture
Vm (U mg−1 )
Km (mM)
Reference
Candida rugosa Candida rugosa Rhizomucor miehei Pseudomonas cepacia
Ethanol/PBS Ethanol/PBS n-Hexane n-Heptane
1.99 3.98 11.72 30
0.76 13 3.03 12
This work [28] [29] [19]
(correlation coefficient (R2 ) of 0.97) with different amount of lipase added. This linear relationship decreased with time because the substrate was gradually consumed. 4. Conclusion
Fig. 5. Kinetics of p-NPP hydrolysis with different amounts (3.56 × 10−3 , 5.34 × 10−3 , 7.13 × 10−3 , 10.69 × 10−3 , 14.25 × 10−3 mg/mL) of lipase and different reactions times (3, 5, 10, 15 and 25 min) under the optimal reaction conditions.
This research centered on design a uniform aqueous-based solution appropriate for the measurement of lipase’s activity. Additionally, the effect of the solution properties on the catalytic activity of C. rugosa lipase was also investigated. To prepare a homogeneous reaction mixture, p-NPP was first dissolved in ethanol at a concentration of 1 mg/mL, and then mixed with PBS at a volume ratio of 1:1. In this aqueous-based solution, the reaction rate was found to be maximum at 45 ◦ C and the pH value of 5.5. In this aqueous solution lipase follows Michealis–Menten saturation kinetics, Vm and Km values are 1.99 U mg−1 and 0.76 mM, respectively. The activation energy of lipase in this solution was 31.71 kJ/mol. This measurement method of lipase catalytic activity in this aqueousbased solution was more convenient and accurate in comparison with previously reported methods. Acknowledgments
different types of lipases in different reaction systems are also listed and compared in Table 1. It was clear that the Km value of lipase in this system (0.76 mM) was lower than Km value in emulsified systems (13 mM) [29]. Vm (maximum reaction rate), which defines the highest possible velocity when all the enzyme is saturated with substrate, reflects the intrinsic characteristics of the enzyme, but may be affected by diffusion constrains [19,30]. The value of this system was also got from the inserted figure in Fig. 4 and the result was 1.99 U/mg.
The authors would like to thank the financial support from the National Natural Science Foundation of China (Grant No. 21274126 and No. 51473143), National “Twelfth Five-Year” Plan for Science & Technology Support of China (No. 2012BAI08B01). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.enzmictec.2015. 01.005.
3.6. Optimal conditions for lipase activity assays On the basis of the experimental results described above, the optimum conditions for lipase catalytic reactions are listed as follows: a buffer/ethanol volume ratio of 1:1, reaction temperature of 45 ◦ C, pH value of 5.5 and p-NPP content of ethanol solution of 1 mg/mL. The process to prepare the reaction system followed the description above. The volume of PBS and p-NPP’s ethanol solution was 1 mL. The amount of p-NPP hydrolysis was measured under conditions with different amounts of enzyme added (7.125 × 10−3 , 10.6875 × 10−3 , 14.25 × 10−3 , 21.375 × 10−3 , 28.5 × 10−3 mg) and different catalytic times (3, 5, 10, 15 and 25 min). As shown in Fig. 5(A), the absorbance was gradually increased with p-NPP hydrolyzed and the substrate was almost completely hydrolyzed after 30 min. It was obvious the more amount of enzyme added the faster the initial hydrolysis rate, and the shorter time required to completely hydrolysis the substrate. Moreover, as the initial content of p-NPP in this reaction solution was 1 mg/mL (2.65 mM) and the liberated p-NP concentration is 2.25 mM after hydrolyzed of 5 min at the lipase concentration of 14.25 × 10−3 mg/mL, so it was nearly 85% of the substrate being hydrolyzed. Fig. 5(B) shows the reaction rates of different lipase addition in different times (3, 5 and 10 min). The initial reaction rates of lipase added amounts of the first 3 min increased nearly linearly
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