- Email: [email protected]
Application of dual-enzyme nanoflower in the epoxidation of alkenes
Process Biochemistry xxx (xxxx) xxx–xxx
Contents lists available at ScienceDirect
Process Biochemistry journal homepage: www.elsevier.com/locate/procbio
Short communication
Application of dual-enzyme nanoflower in the epoxidation of alkenes ⁎
Liu Zhanga, Yuwen Maa, Chunyu Wangb, Zhi Wanga, Xiao Chena, Meixi Lia, Rui Zhaoc, , ⁎ Lei Wanga, a b c
Key Laboratory of Molecular Enzymology and Engineering of Ministry of Education, School of Life Sciences, Jilin University, Changchun 130023, China State Key Laboratory of Supramolecular Structure and Materials, Jilin University, Changchun 130023, China China-Japan Union Hospital of Jilin University, Changchun 130000, China
A R T I C LE I N FO
A B S T R A C T
Keywords: Nanoflower Dual-enzyme Glucose oxidase Lipase Coimmobilization Epoxidation
In this work, a novel organic-inorganic hybrid nanoflower was synthesized by the incorporation of glucose oxidase and lipase and used as the coimmobilized enzyme in the epoxidation of alkenes. Imaging and catalytic performance results suggest that the dual-enzyme nanoflower greatly reduces the diffusion and decomposition of H2O2 and facilitates the mass transfer of glucose oxidase/lipase. Furthermore, the controllable generation of H2O2 and peracid contributes to the good stability of the dual-enzyme nanoflower, which exhibits a higher catalytic performance than the free enzymes in a dual-enzymatic cascade epoxidation. The yield of epoxidation was up to 82% even after ten cycles, further indicating the good reusability of the dual-enzyme nanoflower and its great potential for application in other enzymatic oxidations.
1. Introduction Over the past decade, multi-enzymatic cascade reactions have been become an important and popular topic in biochemical research [1–3]. Such reactions offer considerable advantages, including the regeneration of cofactors, shifting reaction equilibria, and overcoming inhibition effects as well as reducing the reaction time, energy, and material demands of a given process [4–6]. Generally, multi-enzymatic cascade systems must be robust and recyclable for practical application [7]. Coimmobilization is a key technique to increase the operational performance of enzymes and, more importantly, to enhance the catalytic activity of enzymes in multi-enzymatic reactions [8–12]. In addition, the stability and reusability of coimmobilized multi-enzymes have attracted interest from an environmental and economic point of view. In recent decades, the application of glucose oxidase to generate hydrogen peroxide in situ and thereby decrease problems due to inactivation of enzyme by high hydrogen peroxide concentration has been studied in many multi-enzymatic reactions [13,14]. For example, Sheldon’s group has been reported that chloroperoxidase-glucose oxidase system could be applied in the sulfoxidation of thioanisole [15]. This chloroperoxidase-glucose oxidase system has also been used in enantioselective epoxidation of alkenes [16]. Okrasa et al. reported that chiral sulfoxides were synthesized from sulfides using a plant peroxidase from Coprinus cinereus-glucose oxidase bienzymatic system [17]. Recently, a novel dual-enzyme (glucose oxidase and lipase) cascade system for the in situ
⁎
generation of peracid has been applied to the oxidation of various amines and alcohols [18,19]. This system combines the catalytic specificity of glucose oxidase and catalytic promiscuity of lipase to construct a dual-enzyme mediated oxidative process. It also has great potential in some other oxidations, such as Baeyer-Villiger oxidation, Dakin oxidation, sulfonation, and epoxidation [20–25]. Considering the low stability and high cost of free enzymes in biocatalysis, it is highly pertinent to establish a facile and efficient coimmobilization method for this useful dual-enzyme system. In recent years, a simple and versatile immobilization technology to prepare hybrid organic-inorganic nanoflowers composed of various proteins and cupric phosphate has been reported [26]. When an enzyme is encapsulated as the bioactive molecule in the hybrid nanoflower via biomimetic mineralization within the protective exteriors, the enzyme activity and stability is enhanced. Similarly, Ge et al. prepared novel protein-inorganic hybrid nanoflowers for the first time and found that the specific activity of carbonic anhydrase or laccase in the nanoflower was increased dramatically [27]. Wu and his coworkers fabricated lipase-incorporated nanoflower and successfully applied the nanoflower to the enantioselective transesterification of (R,S)-2-pentanol [28]. Jiang et al. reported a facile, economic and green method based on biomimetic mineralization to acquire lipase-inorganic hybrid nanoflower, which was then employed as an economically viable biocatalyst for biodiesel production [29]. Sun et al. reported that the use of the multi-enzyme (glucose oxidase and horseradish peroxidase) co-
Corresponding authors. E-mail addresses: [email protected] (R. Zhao), [email protected] (L. Wang).
https://doi.org/10.1016/j.procbio.2018.08.029 Received 5 July 2018; Received in revised form 13 August 2018; Accepted 25 August 2018 1359-5113/ © 2018 Elsevier Ltd. All rights reserved.
Please cite this article as: Zhang, L., Process Biochemistry, https://doi.org/10.1016/j.procbio.2018.08.029
Process Biochemistry xxx (xxxx) xxx–xxx
L. Zhang et al.
Scheme 1. The process of epoxidation of alkenes mediated by dual-enzyme nanoflower.
protein content of the residual solution was measured by the Lowry method with bovine serium albumin (BSA) as a standard for protein concentration. It’s noteworthy that no protein could be detected in the residual solution after the co-immobilization process which was in accordance with the previous report [28]. Therefore, the enzyme loading efficiency (%) was about 100%. The SEMs of the dual-enzyme nanoflower were obtained from a JSM-6700 F electron microscope (JEOL, Japan). The FTIR spectrums were recorded by Nicolet 5700 FTIR spectrometer (Thermo, USA).
embedded hybrid nanoflower as a colorimetric sensor greatly enhanced the sensitivity of glucose detection [30]. These findings encouraged us to continue trying to coimmobilize glucose oxidase (GOx) and lipase via this facile technique and subsequently apply the dual-enzyme nanoflower in organic oxidations. In this work, we developed a simple method for the preparation of a GOx/lipase nanoflower for the efficient epoxidation of alkenes (Scheme 1). To the best of our knowledge, this is the first time that the incorporation of GOx and lipase in the nanoflower for the epoxidation of alkenes has been reported. In this nanoflower, cupric phosphate was used as the inorganic component, while glucose oxidase from A. niger and lipase B from Candida antarctica (CalB) were adapted as the organic components to construct the dual-enzyme nanoflower. As shown in Scheme 1, when glucose was added into the system, it reacted with O2 and GOx on the dual-enzyme nanoflower to produce H2O2, which was immediately adopted by lipase on the same nanoflower. This was followed by the perhydrolysis of carboxylic ester and resulted in the in situ generation of peracid, which was used in the epoxidation of alkenes. Coimmobilization of this dual-enzyme system can reduce the diffusion resistance of H2O2 to lipase and dramatically accelerate the generation of peracid due to the proximity of the two enzymes that creates a microenvironment for H2O2-rich lipase. More importantly, the controllable generation of H2O2 and peracid by the “feed-on-demand” method in the dual-enzyme nanoflower can avoid enzyme inactivation and improve the stability of the enzyme. The dual-enzyme nanoflower was characterized by scanning electron microscopy (SEM) and Fourier transform infrared spectroscopy (FTIR) and then used in the epoxidation of alkenes, for which reaction conditions were optimized.
2.3. General Procedure for the epoxidation of alkenes mediated by the dualenzyme nanoflower The reaction was performed in a round bottom flask contained alkene (1 mmol), glucose (1.2 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and a mixed solution (5 mL, PB/ EA = 1/9, PB: phosphate buffer (pH 7.0), EA: ethyl acetate). The reaction mixture was stirred at room temperature for 30 h when the oxygen (1 mL/min) was bubbled into the reaction mixture. Then, the mixture was filtered and extracted with dichloromethane. The combined organic phases were dried over Na2SO4 (anhydrous) and concentrated under vacuum, and the resulting residue was purified by flash column chromatography on silica gel with EA/hexane (1/4). The experiments were performed triplicate, and all data were obtained based on the average values. All the isolated products were well characterized by their 1H-NMR spectral analysis. 3. Results and discussion
2. Materials and methods
The dual-enzyme nanoflower was prepared by adding CuSO4 solution to phosphate buffer (pH 7.4) containing GOx and lipase. The system was incubated at 25 °C for three days until a precipitate with porous, flower-like structures appeared. Fig. 1 shows the general morphologies of the dual-enzyme nanoflower imaged by SEM. Fig. 1a reveals that the samples consist of large quantities of flower-like particles with diameters in the range of 20–30 μm. As shown in Fig. 1b, the nanoflower has hierarchical structure with high surface-to-volume ratio. Cu3(PO4)2·3H2O, lipase, GOx, and GOx/lipase nanoflower were characterized by FTIR spectra in the region of 400-4000 cm−1 to certify the enzyme’s presence in the nanoflower (Fig. 2a–d). The vibration bands of PO43− can be seen at 1047 cm−1, 989 cm−1, 628 cm−1 and 559 cm−1 (Fig. 2a). The amide I and II bands of lipase or GOx can be observed at 1646 cm−1 and 1533 cm−1 (Fig. 2b and c), respectively. The spectra of GOx/lipase nanoflower (Fig. 2d) indicates that the enzymes were incorporated in the nanoflower successfully. Subsequently, the loading efficiency of the dual-enzyme nanoflower was investigated using 100 mg lipase and 100 mg GOx. After the immobilization process was performed, the protein content in the pooled suspension and washing solution was measured. Since no protein was detected in the above-mentioned solution, the loading efficiency (%)
2.1. Materials Glucose oxidase from A. Niger (GOx) and Candida antarctica lipase B (CalB) were purchased from Sigma (Beijing, China). These enzymes were used after lyophilization for the immobilization without further purification. Glucose and alkenes used in this study were purchased from J&K Scientific (Beijing, China). All the other chemical reagents were purchased from Shanghai Chemical Reagent Company (Shanghai, China). All the commercially available reagents and solvents were used without further purification. NMR spectra were recorded on an Inova 500 (500 MHz) spectrometer. 2.2. Preparation of the dual-enzyme nanoflower CuSO4 solution (120 mM) was added to the phosphate buffer (pH 7.4, 100 mL) containing GOx (glucose oxidase from A. niger, 100 mg) and lipase (lipase B from Candida antarctica, 100 mg). The system was incubated at 25 °C for 72 h. Then a precipitate with porous, flower-like structures appeared. The prepared dual-enzyme nanoflowers were separated by centrifugation and dried overnight under vacuum. The 2
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Fig. 1. Scanning electron microscopy (SEM) images of the GOx/lipase nanoflower.
nanoflower has great potential for application in the epoxidation of styrene. Generally, phosphate buffer (PB) is the common solvent for GOX catalyzed reactions [31]. And ethyl acetate (EA) can be adopted as the substrate and reaction medium for the lipase-mediated oxidation [32]. Thus, a biphasic reaction medium (PB and EA) is needed for this reaction. In this work, the effect of the volume ratio (PB/EA) was investigated when the amount of glucose was fixed at 1.2 mmol, and the results are shown in Table 2 (Entry 1–5). It was found that the yield of styrene oxide was increased by increasing the volume of EA, and a satisfactory yield (89%) was obtained at the volume ratio (PB/EA) of 1/ 9 for this reaction. The increasing of the volume of EA might enhance the generation rate of peracid. Besides, a better solubility of substrate offered by EA could accelerate the reaction in some extent. However, the activity of GOx might be decreased in a low content of PB, and then decreased the catalytic performance of dual-enzyme nanoflower. Therefore, a volume ratio (PB/EA) of 1/9 was used for all the subsequent experiments. The effect of the amount of glucose was also studied in this work. It could be found in Table 2 (Entry 4, 6–8) that 1.2 mmol glucose was the ideal amount for this reaction. Further increasing the amount of glucose did not improve the yield of epoxidation obviously. Considering that enzyme reusability is an important parameter to evaluate an immobilized enzyme in practical application [33], we investigated the reusability of the GOx/lipase nanoflower (Fig. 3). It was found that 82% epoxidation yield of styrene could be obtained even after ten cycles. This result reveals that the dual-enzyme nanoflower possesses excellent reusability, which may facilitate the epoxidation of styrene during a continuous operation. However, the slight decrease in epoxidation yield might be due to the leakage of the enzyme from the nanoflower and slight inactivation of the enzyme in the reaction cycles [34,35]. Under the optimum conditions, the time course of this epoxidation mediated by dual-enzyme nanoflower and free dual-enzyme was studied. As shown in Fig. 4, the reaction reached its equilibrium in
Fig. 2. FTIR spectra of Cu3(PO4)2⋅3H2O, lipase, GOx, and GOx/lipase nanoflower.
was about 100%. Since 1.1 g dual-enzyme nanoflower was obtained, the enzyme loading amount in the nanoflower was about 180 mg enzyme/g nanoflower. To illustrate the advantage of the GOx/lipase nanoflower, we compared its catalytic performance in the epoxidation of styrene with that of a free dual-enzyme. As shown in Table 1, the catalytic performance of the dual-enzyme nanoflower was much higher than that of the free dual-enzyme. Moreover, the performance of the GOx/lipase nanoflower was also 50% higher than that of a mixture of GOx nanoflower and lipase nanoflower with a mass ratio of 1:1 (protein content, GOx: 9 mg, lipase: 9 mg). All the results suggest that this coimmobilized method largely reduced the diffusion resistance and concentration of H2O2, thereby greatly avoiding enzyme inactivation, improving the generation rate of peracid, and increasing the yield of styrene oxide. Furthermore, the coimmobilization could preserve the active conformation of the protein and keep the activities of the two enzymes in the nanoflower. These results further indicate that this dual-enzyme
Table 2 Effects of reaction medium and amount of glucose on the epoxidation. Styrene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and glucose were added in reaction medium (5 mL), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h.
Table 1 Catalytic performance of enzyme nanoflowers and free enzymes on the epoxidation. Styrene (1 mmol), catalyst (protein content: 18 mg, lipase/GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (phosphate buffer (PB, pH 7.0, 0.5 mL) and ethyl acetate (EA, 4.5 mL)), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h. Entry
Catalyst
Yield (%)
1 2
Free GOx and lipase Free GOx and lipase in the presence of copper phosphate crystals GOx nanoflower and lipase nanoflower GOx/lipase nanoflower
51 ± 1.6 53 ± 1.2
3 4
65 ± 1.1 89 ± 0.8
3
Entry
Volume ratio (PB/EA)
Amount of glucose (mmol)
Yield (%)
1 2 3 4 5 6 7 8
1/1 1/3 1/6 1/9 1/12 1/9 1/9 1/9
1.2 1.2 1.2 1.2 1.2 1 1.4 1.6
63 75 81 89 83 78 91 91
± ± ± ± ± ± ± ±
2.4 1.9 1.5 0.8 1.2 2.4 0.7 0.3
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Table 3 Epoxidation of alkenes mediated by dual-enzyme nanoflower. Alkene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (5 mL, PB/EA = 1/9), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h. Entry
Fig. 3. The reusability of the dual-enzyme nanoflower on the epoxidation. Styrene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/ GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (5 mL, PB/EA = 1/9), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h.
Alkene
Yield (%)
1
89 ± 0.8
2
83 ± 1.6
3
93 ± 0.5
4
77 ± 2.7
5
94 ± 0.9
6
96 ± 0.4
enzyme nanoflower exhibits a much higher catalytic performance in the epoxidation of alkenes and excellent reusability, which was observed in this reaction system. Moreover, this study provides a new case of multienzyme coimmobilized organic-inorganic hybrid nanoflowers for multistep cascade enzymatic reactions. Acknowledgements We gratefully acknowledge the Foundation of Changchun BC&HC Pharmaceutical Technology Co., Ltd (no. 3R117W391465). Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.procbio.2018.08.029. References [1] E. Ricca, B. Brucher, J.H. Schrittwieser, Multi-enzymatic cascade reactions: overview and perspectives, Adv. Synth. Catal. 353 (2011) 2239–2262. [2] F. Lopez-Gallego, C. Schmidt-Dannert, Multi-enzymatic synthesis, Curr. Opin. Chem. Biol. 14 (2010) 174–183. [3] R. Xue, J.M. Woodley, Process technology for multi-enzymatic reaction systems, Bioresour. Technol. Rep. 115 (2012) 183–195. [4] S. Schoffelen, J.C. van Hest, Multi-enzyme systems: bringing enzymes together in vitro, Soft Matter 8 (2012) 1736–1746. [5] S. Schoffelen, J.C. van Hest, Chemical approaches for the construction of multienzyme reaction systems, Curr. Opin. Struct. Biol. 23 (2013) 613–621. [6] W.D. Fessner, Systems biocatalysis: development and engineering of cell-free “artificial metabolisms” for preparative multi-enzymatic synthesis, New Biotechnol. 32 (2015) 658–664. [7] J. Rocha-Martín, B.D.L. Rivas, R. Muñoz, J.M. Guisán, F. López-Gallego, Rational co-immobilization of bi-enzyme cascades on porous supports and their applications in bio-redox reactions with in situ recycling of soluble cofactors, ChemCatChem 4 (2012) 1279–1288. [8] S. Fornera, P. Kuhn, D. Lombardi, A.D. Schlüter, P.S. Dittrich, P. Walde, Sequential immobilization of enzymes in microfluidic channels for cascade reactions, ChemPlusChem 77 (2012) 98–101. [9] S. Talekar, A. Pandharbale, M. Ladole, S. Nadar, M. Mulla, K. Japhalekar, P. Kishori, D. Arage, Carrier free co-immobilization of alpha amylase, glucoamylase and pullulanase as combined cross-linked enzyme aggregates (combi-CLEAs): a tri-enzyme biocatalyst with one pot starch hydrolytic activity, Bioresour. Technol. 147 (2013) 269–275. [10] J. Chung, E.T. Hwang, J.H. Kim, B.C. Kim, M.B. Gu, Modular multi-enzyme cascade process using highly stabilized enzyme microbeads, Green Chem. 16 (2014) 1163–1167. [11] I. Aranaz, V. Ramos, S. De La Escalera, A. Heras, Co-immobilization of D-hydantoinase and D-carboamylase on chitin: application to the synthesis of p-hydroxyphenylglycine, Biocatal. Biotransfor. 21 (2003) 349–356. [12] F. Zhao, H. Li, Y. Jiang, X. Wang, X. Mu, Co-immobilization of multi-enzyme on control-reduced graphene oxide by non-covalent bonds: an artificial biocatalytic system for the one-pot production of gluconic acid from starch, Green Chem. 16
Fig. 4. The time course of the epoxidation mediated by dual-enzyme nanoflower and free dual-enzyme. Styrene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (5 mL, PB/EA = 1/9), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature.
approximately 30 h with an epoxidation yield of 89% when dual-enzyme nanoflower was used in this epoxidation. However, the catalytic performance of free dual-enzyme was much lower and only 58% yield could be obtained even after 40 h. This observation indicates that the coimmobilization can obviously enhance dual-enzyme catalytic performance and increase the final epoxidation yield. This novel dual-enzyme nanoflower was also used for the epoxidation of other alkenes. Table 3 shows that all selected alkenes could be epoxidized to the corresponding products with high yields (77–96%). The good generality of the dual-enzyme nanoflower in the epoxidation demonstrates that it has great potential for practical oxidative processes. 4. Conclusion In this study, a novel dual-enzyme nanoflower was successfully prepared for the coimmobilization of GOx and lipase and used in the epoxidation of alkenes. Compared with the free dual-enzyme, the dual4
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[25] M. Svedendahl, P. Carlqvist, C. Branneby, O. Allnér, A. Frise, K. Hult, P. Berglund, T. Brinck, Direct epoxidation in Candida antarctica lipase B studied by experiment and theory, ChemBioChem 9 (2008) 2443–2451. [26] X. Wu, M. Hou, J. Ge, Metal-organic frameworks and inorganic nanoflowers: a type of emerging inorganic crystal nanocarrier for enzyme immobilization, Catal. Sci. Technol. 5 (2015) 5077–5085. [27] J. Ge, J. Lei, R.N. Zare, Protein-inorganic hybrid nanoflowers, Nat. Nanotechnol. 7 (2012) 428. [28] Z. Wu, H. Li, X. Zhu, S. Li, Z. Wang, L. Wang, Z. Li, G. Chen, Using laccases in the nanoflower to synthesize Viniferin, Catalysts 7 (2017) 188. [29] W. Jiang, X. Wang, J. Yang, H. Han, Q. Li, J. Tang, Lipase-inorganic hybrid nanoflower constructed through biomimetic mineralization: a new support for biodiesel synthesis, J. Colloid Interface Sci. 514 (2018) 102–107. [30] J. Sun, J. Ge, W. Liu, M. Lan, H. Zhang, P. Wang, Y. Wang, Z. Niu, Multi-enzyme coembedded organic-inorganic hybrid nanoflowers: synthesis and application as a colorimetric sensor, Nanoscale 6 (2014) 255–262. [31] D. Jung, C. Streb, M. Hartmann, Oxidation of indole using chloroperoxidase and glucose oxidase immobilized on SBA-15 as tandem biocatalyst, Microporous Mesoporous Mater. 113 (2008) 523–529. [32] F. Yang, Z. Wang, X. Zhang, L. Jiang, Y. Li, L. Wang, A green chemoenzymatic process for the synthesis of azoxybenzenes, ChemCatChem 7 (2015) 3450–3453. [33] J.C.S.D. Santos, O. Barbosa, C. Ortiz, A. Berenguer-Murcia, R.C. Rodrigues, R. Fernandez-Lafuente, Importance of the support properties for immobilization or purification of enzymes, ChemCatChem 7 (2015) 2413–2432. [34] L. Fernandez-Lopez, J.J. Virgen-OrtÍz, S.G. Pedrero, N. Lopez-Carrobles, B.C. Gorines, C. Otero, R. Fernandez-Lafuente, Optimization of the coating of octylCALB with ionic polymers to improve stability and decrease enzyme leakage, Biocatal. Biotransfor. 36 (2018) 47–56. [35] J.J. Virgen-Ortíz, S. Peirce, V.G. Tacias-Pascacio, V. Cortes-Corberan, A. Marzocchella, M.E. Russo, R. Fernandez-Lafuente, Reuse of anion exchangers as supports for enzyme immobilization: reinforcement of the enzyme-support multiinteraction after enzyme inactivation, Process Biochem. 51 (2016) 1391–1396.
(2014) 2558–2565. [13] F. van de Velde, N.D. Lourenço, M. Bakker, F. van Rantwijk, R.A. Sheldon, Improved operational stability of peroxidases by coimmobilization with glucose oxidase, Biotechnol. Bioeng. 69 (2000) 286–291. [14] F. van de Velde, F. van Rantwijk, R.A. Sheldon, Improving the catalytic performance of peroxidases in organic synthesis, Trends Biotechnol. 19 (2001) 73–80. [15] P.C. Pereira, I.W. Arends, R.A. Sheldon, Optimizing the chloroperoxidase-glucose oxidase system: the effect of glucose oxidase on activity and enantioselectivity, Process Biochem. 50 (2015) 746–751. [16] R. Narayanan, G. Zhu, P. Wang, Stabilization of interface-binding chloroperoxidase for interfacial biotransformation, J. Biotechnol. 128 (2007) 86–92. [17] K. Okrasa, E. Guibé-Jampel, M. Therisod, Tandem peroxidase-glucose oxidase catalysed enantioselective sulfoxidation of thioanisoles, J. Chem. Soc. Perkin Trans. 1 (7) (2000) 1077–1079. [18] F. Yang, X. Zhang, F. Li, Z. Wang, L. Wang, A lipase-glucose oxidase system for the efficient oxidation of N-heteroaromatic compounds and tertiary amines, Green Chem. 18 (2016) 3518–3521. [19] D. Thiel, D. Doknić, J. Deska, Enzymatic aerobic ring rearrangement of optically active furylcarbinols, Nat. Commun. 5 (2014) 5278. [20] F. Blume, P. Sprengart, J. Deska, Lipase-induced oxidative furan rearrangements, Synlett 29 (2018) 1293–1296. [21] Z. Wang, X. Chen, C. Wang, L. Zhang, F. Li, W. Zhang, P. Chen, L. Wang, A mild and efficient Dakin reaction mediated by lipase, Green Chem. Lett. Rev. 10 (2017) 269–273. [22] A.J. Kotlewska, F. van Rantwijk, R.A. Sheldon, I.W. Arends, Epoxidation and Baeyer-Villiger oxidation using hydrogen peroxide and a lipase dissolved in ionic liquids, Green Chem. 13 (2011) 2154–2160. [23] M.Y. Ríos, E. Salazar, H.F. Olivo, Baeyer-Villiger oxidation of substituted cyclohexanones via lipase-mediated perhydrolysis utilizing urea-hydrogen peroxide in ethyl acetate, Green Chem. 9 (2007) 459–462. [24] E.G. Ankudey, H.F. Olivo, T.L. Peeples, Lipase-mediated epoxidation utilizing urea–hydrogen peroxide in ethyl acetate, Green Chem. 8 (2006) 923–926.
5
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Short communication
Application of dual-enzyme nanoflower in the epoxidation of alkenes ⁎
Liu Zhanga, Yuwen Maa, Chunyu Wangb, Zhi Wanga, Xiao Chena, Meixi Lia, Rui Zhaoc, , ⁎ Lei Wanga, a b c
Key Laboratory of Molecular Enzymology and Engineering of Ministry of Education, School of Life Sciences, Jilin University, Changchun 130023, China State Key Laboratory of Supramolecular Structure and Materials, Jilin University, Changchun 130023, China China-Japan Union Hospital of Jilin University, Changchun 130000, China
A R T I C LE I N FO
A B S T R A C T
Keywords: Nanoflower Dual-enzyme Glucose oxidase Lipase Coimmobilization Epoxidation
In this work, a novel organic-inorganic hybrid nanoflower was synthesized by the incorporation of glucose oxidase and lipase and used as the coimmobilized enzyme in the epoxidation of alkenes. Imaging and catalytic performance results suggest that the dual-enzyme nanoflower greatly reduces the diffusion and decomposition of H2O2 and facilitates the mass transfer of glucose oxidase/lipase. Furthermore, the controllable generation of H2O2 and peracid contributes to the good stability of the dual-enzyme nanoflower, which exhibits a higher catalytic performance than the free enzymes in a dual-enzymatic cascade epoxidation. The yield of epoxidation was up to 82% even after ten cycles, further indicating the good reusability of the dual-enzyme nanoflower and its great potential for application in other enzymatic oxidations.
1. Introduction Over the past decade, multi-enzymatic cascade reactions have been become an important and popular topic in biochemical research [1–3]. Such reactions offer considerable advantages, including the regeneration of cofactors, shifting reaction equilibria, and overcoming inhibition effects as well as reducing the reaction time, energy, and material demands of a given process [4–6]. Generally, multi-enzymatic cascade systems must be robust and recyclable for practical application [7]. Coimmobilization is a key technique to increase the operational performance of enzymes and, more importantly, to enhance the catalytic activity of enzymes in multi-enzymatic reactions [8–12]. In addition, the stability and reusability of coimmobilized multi-enzymes have attracted interest from an environmental and economic point of view. In recent decades, the application of glucose oxidase to generate hydrogen peroxide in situ and thereby decrease problems due to inactivation of enzyme by high hydrogen peroxide concentration has been studied in many multi-enzymatic reactions [13,14]. For example, Sheldon’s group has been reported that chloroperoxidase-glucose oxidase system could be applied in the sulfoxidation of thioanisole [15]. This chloroperoxidase-glucose oxidase system has also been used in enantioselective epoxidation of alkenes [16]. Okrasa et al. reported that chiral sulfoxides were synthesized from sulfides using a plant peroxidase from Coprinus cinereus-glucose oxidase bienzymatic system [17]. Recently, a novel dual-enzyme (glucose oxidase and lipase) cascade system for the in situ
⁎
generation of peracid has been applied to the oxidation of various amines and alcohols [18,19]. This system combines the catalytic specificity of glucose oxidase and catalytic promiscuity of lipase to construct a dual-enzyme mediated oxidative process. It also has great potential in some other oxidations, such as Baeyer-Villiger oxidation, Dakin oxidation, sulfonation, and epoxidation [20–25]. Considering the low stability and high cost of free enzymes in biocatalysis, it is highly pertinent to establish a facile and efficient coimmobilization method for this useful dual-enzyme system. In recent years, a simple and versatile immobilization technology to prepare hybrid organic-inorganic nanoflowers composed of various proteins and cupric phosphate has been reported [26]. When an enzyme is encapsulated as the bioactive molecule in the hybrid nanoflower via biomimetic mineralization within the protective exteriors, the enzyme activity and stability is enhanced. Similarly, Ge et al. prepared novel protein-inorganic hybrid nanoflowers for the first time and found that the specific activity of carbonic anhydrase or laccase in the nanoflower was increased dramatically [27]. Wu and his coworkers fabricated lipase-incorporated nanoflower and successfully applied the nanoflower to the enantioselective transesterification of (R,S)-2-pentanol [28]. Jiang et al. reported a facile, economic and green method based on biomimetic mineralization to acquire lipase-inorganic hybrid nanoflower, which was then employed as an economically viable biocatalyst for biodiesel production [29]. Sun et al. reported that the use of the multi-enzyme (glucose oxidase and horseradish peroxidase) co-
Corresponding authors. E-mail addresses: [email protected] (R. Zhao), [email protected] (L. Wang).
https://doi.org/10.1016/j.procbio.2018.08.029 Received 5 July 2018; Received in revised form 13 August 2018; Accepted 25 August 2018 1359-5113/ © 2018 Elsevier Ltd. All rights reserved.
Please cite this article as: Zhang, L., Process Biochemistry, https://doi.org/10.1016/j.procbio.2018.08.029
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Scheme 1. The process of epoxidation of alkenes mediated by dual-enzyme nanoflower.
protein content of the residual solution was measured by the Lowry method with bovine serium albumin (BSA) as a standard for protein concentration. It’s noteworthy that no protein could be detected in the residual solution after the co-immobilization process which was in accordance with the previous report [28]. Therefore, the enzyme loading efficiency (%) was about 100%. The SEMs of the dual-enzyme nanoflower were obtained from a JSM-6700 F electron microscope (JEOL, Japan). The FTIR spectrums were recorded by Nicolet 5700 FTIR spectrometer (Thermo, USA).
embedded hybrid nanoflower as a colorimetric sensor greatly enhanced the sensitivity of glucose detection [30]. These findings encouraged us to continue trying to coimmobilize glucose oxidase (GOx) and lipase via this facile technique and subsequently apply the dual-enzyme nanoflower in organic oxidations. In this work, we developed a simple method for the preparation of a GOx/lipase nanoflower for the efficient epoxidation of alkenes (Scheme 1). To the best of our knowledge, this is the first time that the incorporation of GOx and lipase in the nanoflower for the epoxidation of alkenes has been reported. In this nanoflower, cupric phosphate was used as the inorganic component, while glucose oxidase from A. niger and lipase B from Candida antarctica (CalB) were adapted as the organic components to construct the dual-enzyme nanoflower. As shown in Scheme 1, when glucose was added into the system, it reacted with O2 and GOx on the dual-enzyme nanoflower to produce H2O2, which was immediately adopted by lipase on the same nanoflower. This was followed by the perhydrolysis of carboxylic ester and resulted in the in situ generation of peracid, which was used in the epoxidation of alkenes. Coimmobilization of this dual-enzyme system can reduce the diffusion resistance of H2O2 to lipase and dramatically accelerate the generation of peracid due to the proximity of the two enzymes that creates a microenvironment for H2O2-rich lipase. More importantly, the controllable generation of H2O2 and peracid by the “feed-on-demand” method in the dual-enzyme nanoflower can avoid enzyme inactivation and improve the stability of the enzyme. The dual-enzyme nanoflower was characterized by scanning electron microscopy (SEM) and Fourier transform infrared spectroscopy (FTIR) and then used in the epoxidation of alkenes, for which reaction conditions were optimized.
2.3. General Procedure for the epoxidation of alkenes mediated by the dualenzyme nanoflower The reaction was performed in a round bottom flask contained alkene (1 mmol), glucose (1.2 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and a mixed solution (5 mL, PB/ EA = 1/9, PB: phosphate buffer (pH 7.0), EA: ethyl acetate). The reaction mixture was stirred at room temperature for 30 h when the oxygen (1 mL/min) was bubbled into the reaction mixture. Then, the mixture was filtered and extracted with dichloromethane. The combined organic phases were dried over Na2SO4 (anhydrous) and concentrated under vacuum, and the resulting residue was purified by flash column chromatography on silica gel with EA/hexane (1/4). The experiments were performed triplicate, and all data were obtained based on the average values. All the isolated products were well characterized by their 1H-NMR spectral analysis. 3. Results and discussion
2. Materials and methods
The dual-enzyme nanoflower was prepared by adding CuSO4 solution to phosphate buffer (pH 7.4) containing GOx and lipase. The system was incubated at 25 °C for three days until a precipitate with porous, flower-like structures appeared. Fig. 1 shows the general morphologies of the dual-enzyme nanoflower imaged by SEM. Fig. 1a reveals that the samples consist of large quantities of flower-like particles with diameters in the range of 20–30 μm. As shown in Fig. 1b, the nanoflower has hierarchical structure with high surface-to-volume ratio. Cu3(PO4)2·3H2O, lipase, GOx, and GOx/lipase nanoflower were characterized by FTIR spectra in the region of 400-4000 cm−1 to certify the enzyme’s presence in the nanoflower (Fig. 2a–d). The vibration bands of PO43− can be seen at 1047 cm−1, 989 cm−1, 628 cm−1 and 559 cm−1 (Fig. 2a). The amide I and II bands of lipase or GOx can be observed at 1646 cm−1 and 1533 cm−1 (Fig. 2b and c), respectively. The spectra of GOx/lipase nanoflower (Fig. 2d) indicates that the enzymes were incorporated in the nanoflower successfully. Subsequently, the loading efficiency of the dual-enzyme nanoflower was investigated using 100 mg lipase and 100 mg GOx. After the immobilization process was performed, the protein content in the pooled suspension and washing solution was measured. Since no protein was detected in the above-mentioned solution, the loading efficiency (%)
2.1. Materials Glucose oxidase from A. Niger (GOx) and Candida antarctica lipase B (CalB) were purchased from Sigma (Beijing, China). These enzymes were used after lyophilization for the immobilization without further purification. Glucose and alkenes used in this study were purchased from J&K Scientific (Beijing, China). All the other chemical reagents were purchased from Shanghai Chemical Reagent Company (Shanghai, China). All the commercially available reagents and solvents were used without further purification. NMR spectra were recorded on an Inova 500 (500 MHz) spectrometer. 2.2. Preparation of the dual-enzyme nanoflower CuSO4 solution (120 mM) was added to the phosphate buffer (pH 7.4, 100 mL) containing GOx (glucose oxidase from A. niger, 100 mg) and lipase (lipase B from Candida antarctica, 100 mg). The system was incubated at 25 °C for 72 h. Then a precipitate with porous, flower-like structures appeared. The prepared dual-enzyme nanoflowers were separated by centrifugation and dried overnight under vacuum. The 2
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Fig. 1. Scanning electron microscopy (SEM) images of the GOx/lipase nanoflower.
nanoflower has great potential for application in the epoxidation of styrene. Generally, phosphate buffer (PB) is the common solvent for GOX catalyzed reactions [31]. And ethyl acetate (EA) can be adopted as the substrate and reaction medium for the lipase-mediated oxidation [32]. Thus, a biphasic reaction medium (PB and EA) is needed for this reaction. In this work, the effect of the volume ratio (PB/EA) was investigated when the amount of glucose was fixed at 1.2 mmol, and the results are shown in Table 2 (Entry 1–5). It was found that the yield of styrene oxide was increased by increasing the volume of EA, and a satisfactory yield (89%) was obtained at the volume ratio (PB/EA) of 1/ 9 for this reaction. The increasing of the volume of EA might enhance the generation rate of peracid. Besides, a better solubility of substrate offered by EA could accelerate the reaction in some extent. However, the activity of GOx might be decreased in a low content of PB, and then decreased the catalytic performance of dual-enzyme nanoflower. Therefore, a volume ratio (PB/EA) of 1/9 was used for all the subsequent experiments. The effect of the amount of glucose was also studied in this work. It could be found in Table 2 (Entry 4, 6–8) that 1.2 mmol glucose was the ideal amount for this reaction. Further increasing the amount of glucose did not improve the yield of epoxidation obviously. Considering that enzyme reusability is an important parameter to evaluate an immobilized enzyme in practical application [33], we investigated the reusability of the GOx/lipase nanoflower (Fig. 3). It was found that 82% epoxidation yield of styrene could be obtained even after ten cycles. This result reveals that the dual-enzyme nanoflower possesses excellent reusability, which may facilitate the epoxidation of styrene during a continuous operation. However, the slight decrease in epoxidation yield might be due to the leakage of the enzyme from the nanoflower and slight inactivation of the enzyme in the reaction cycles [34,35]. Under the optimum conditions, the time course of this epoxidation mediated by dual-enzyme nanoflower and free dual-enzyme was studied. As shown in Fig. 4, the reaction reached its equilibrium in
Fig. 2. FTIR spectra of Cu3(PO4)2⋅3H2O, lipase, GOx, and GOx/lipase nanoflower.
was about 100%. Since 1.1 g dual-enzyme nanoflower was obtained, the enzyme loading amount in the nanoflower was about 180 mg enzyme/g nanoflower. To illustrate the advantage of the GOx/lipase nanoflower, we compared its catalytic performance in the epoxidation of styrene with that of a free dual-enzyme. As shown in Table 1, the catalytic performance of the dual-enzyme nanoflower was much higher than that of the free dual-enzyme. Moreover, the performance of the GOx/lipase nanoflower was also 50% higher than that of a mixture of GOx nanoflower and lipase nanoflower with a mass ratio of 1:1 (protein content, GOx: 9 mg, lipase: 9 mg). All the results suggest that this coimmobilized method largely reduced the diffusion resistance and concentration of H2O2, thereby greatly avoiding enzyme inactivation, improving the generation rate of peracid, and increasing the yield of styrene oxide. Furthermore, the coimmobilization could preserve the active conformation of the protein and keep the activities of the two enzymes in the nanoflower. These results further indicate that this dual-enzyme
Table 2 Effects of reaction medium and amount of glucose on the epoxidation. Styrene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and glucose were added in reaction medium (5 mL), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h.
Table 1 Catalytic performance of enzyme nanoflowers and free enzymes on the epoxidation. Styrene (1 mmol), catalyst (protein content: 18 mg, lipase/GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (phosphate buffer (PB, pH 7.0, 0.5 mL) and ethyl acetate (EA, 4.5 mL)), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h. Entry
Catalyst
Yield (%)
1 2
Free GOx and lipase Free GOx and lipase in the presence of copper phosphate crystals GOx nanoflower and lipase nanoflower GOx/lipase nanoflower
51 ± 1.6 53 ± 1.2
3 4
65 ± 1.1 89 ± 0.8
3
Entry
Volume ratio (PB/EA)
Amount of glucose (mmol)
Yield (%)
1 2 3 4 5 6 7 8
1/1 1/3 1/6 1/9 1/12 1/9 1/9 1/9
1.2 1.2 1.2 1.2 1.2 1 1.4 1.6
63 75 81 89 83 78 91 91
± ± ± ± ± ± ± ±
2.4 1.9 1.5 0.8 1.2 2.4 0.7 0.3
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Table 3 Epoxidation of alkenes mediated by dual-enzyme nanoflower. Alkene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (5 mL, PB/EA = 1/9), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h. Entry
Fig. 3. The reusability of the dual-enzyme nanoflower on the epoxidation. Styrene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/ GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (5 mL, PB/EA = 1/9), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature for 30 h.
Alkene
Yield (%)
1
89 ± 0.8
2
83 ± 1.6
3
93 ± 0.5
4
77 ± 2.7
5
94 ± 0.9
6
96 ± 0.4
enzyme nanoflower exhibits a much higher catalytic performance in the epoxidation of alkenes and excellent reusability, which was observed in this reaction system. Moreover, this study provides a new case of multienzyme coimmobilized organic-inorganic hybrid nanoflowers for multistep cascade enzymatic reactions. Acknowledgements We gratefully acknowledge the Foundation of Changchun BC&HC Pharmaceutical Technology Co., Ltd (no. 3R117W391465). Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.procbio.2018.08.029. References [1] E. Ricca, B. Brucher, J.H. Schrittwieser, Multi-enzymatic cascade reactions: overview and perspectives, Adv. Synth. Catal. 353 (2011) 2239–2262. [2] F. Lopez-Gallego, C. Schmidt-Dannert, Multi-enzymatic synthesis, Curr. Opin. Chem. Biol. 14 (2010) 174–183. [3] R. Xue, J.M. Woodley, Process technology for multi-enzymatic reaction systems, Bioresour. Technol. Rep. 115 (2012) 183–195. [4] S. Schoffelen, J.C. van Hest, Multi-enzyme systems: bringing enzymes together in vitro, Soft Matter 8 (2012) 1736–1746. [5] S. Schoffelen, J.C. van Hest, Chemical approaches for the construction of multienzyme reaction systems, Curr. Opin. Struct. Biol. 23 (2013) 613–621. [6] W.D. Fessner, Systems biocatalysis: development and engineering of cell-free “artificial metabolisms” for preparative multi-enzymatic synthesis, New Biotechnol. 32 (2015) 658–664. [7] J. Rocha-Martín, B.D.L. Rivas, R. Muñoz, J.M. Guisán, F. López-Gallego, Rational co-immobilization of bi-enzyme cascades on porous supports and their applications in bio-redox reactions with in situ recycling of soluble cofactors, ChemCatChem 4 (2012) 1279–1288. [8] S. Fornera, P. Kuhn, D. Lombardi, A.D. Schlüter, P.S. Dittrich, P. Walde, Sequential immobilization of enzymes in microfluidic channels for cascade reactions, ChemPlusChem 77 (2012) 98–101. [9] S. Talekar, A. Pandharbale, M. Ladole, S. Nadar, M. Mulla, K. Japhalekar, P. Kishori, D. Arage, Carrier free co-immobilization of alpha amylase, glucoamylase and pullulanase as combined cross-linked enzyme aggregates (combi-CLEAs): a tri-enzyme biocatalyst with one pot starch hydrolytic activity, Bioresour. Technol. 147 (2013) 269–275. [10] J. Chung, E.T. Hwang, J.H. Kim, B.C. Kim, M.B. Gu, Modular multi-enzyme cascade process using highly stabilized enzyme microbeads, Green Chem. 16 (2014) 1163–1167. [11] I. Aranaz, V. Ramos, S. De La Escalera, A. Heras, Co-immobilization of D-hydantoinase and D-carboamylase on chitin: application to the synthesis of p-hydroxyphenylglycine, Biocatal. Biotransfor. 21 (2003) 349–356. [12] F. Zhao, H. Li, Y. Jiang, X. Wang, X. Mu, Co-immobilization of multi-enzyme on control-reduced graphene oxide by non-covalent bonds: an artificial biocatalytic system for the one-pot production of gluconic acid from starch, Green Chem. 16
Fig. 4. The time course of the epoxidation mediated by dual-enzyme nanoflower and free dual-enzyme. Styrene (1 mmol), dual-enzyme nanoflower (protein content: 18 mg, lipase/GOx = 1/1) and glucose (1.2 mmol) were added in reaction medium (5 mL, PB/EA = 1/9), and oxygen (1 mL/min) was bubbled into the reaction mixture. The reaction was performed at room temperature.
approximately 30 h with an epoxidation yield of 89% when dual-enzyme nanoflower was used in this epoxidation. However, the catalytic performance of free dual-enzyme was much lower and only 58% yield could be obtained even after 40 h. This observation indicates that the coimmobilization can obviously enhance dual-enzyme catalytic performance and increase the final epoxidation yield. This novel dual-enzyme nanoflower was also used for the epoxidation of other alkenes. Table 3 shows that all selected alkenes could be epoxidized to the corresponding products with high yields (77–96%). The good generality of the dual-enzyme nanoflower in the epoxidation demonstrates that it has great potential for practical oxidative processes. 4. Conclusion In this study, a novel dual-enzyme nanoflower was successfully prepared for the coimmobilization of GOx and lipase and used in the epoxidation of alkenes. Compared with the free dual-enzyme, the dual4
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