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Nanostructured Biocatalysts
2.67
Nanostructured Biocatalysts
J Ge, D Lu, M Yan, and Z Liu, Tsinghua University, Beijing, China © 2011 Elsevier B.V. All rights reserved.
2.67.1 2.67.2 2.67.3 2.67.4 2.67.4.1 2.67.4.2 2.67.4.2.1 2.67.4.2.2 2.67.4.2.3 2.67.5 2.67.5.1 2.67.5.2 2.67.5.3 2.67.6 2.67.6.1 2.67.6.2 2.67.6.3 2.67.7 References
Introduction Nonaqueous Enzymatic Catalysis Enzymes in Nanostructures Preparation of Enzyme Nanogels Synthetic Procedure Optimization of the Synthetic Reactions Used for Lipase Nanogel Preparation The pH of the acryloylation reaction Monomers for in situ polymerization Monomer concentration in the polymerization reaction Molecular Fundamentals of Enzyme Nanogels Assembly of Monomers around the Enzyme in Aqueous Solution Enzyme Stabilization via the Polymer Network Experimental Validation of Enhanced Enzyme Stability Potential Applications of Enzyme Nanogels as Biocatalysts Lipase Nanogel-Catalyzed Synthesis of Biodiesel Lipase Nanogel-Catalyzed Synthesis of a Dextran-Based Surfactant Lipase Nanogel-Catalyzed Synthesis of Polyester Summary
Glossary enzymatic catalysis The chemical reactions catalyzed by an enzyme that naturally catalyzes reactions in a biological system. enzyme stabilization The physical or chemical methods to enhance enzyme stability under adverse conditions such as high temperature or the presence of denaturing solvents. enzyme modification The chemical methods that introduce additional structure to an enzyme or to enhance enzyme stability and/or activity.
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enzyme nanogel The transparent polymer matrix with diameter below 100 nm, which is obtained by in situ polymerization from the enzyme surface. nanostructured enzyme catalyst The materials below 100 nm in dimension(s) such as nanobeads and nanofibers, which display the activity of the incorporated enzyme.
2.67.1 Introduction Enzyme stabilization is of fundamental importance for extending the scope of enzymatic catalysis to nonaqueous media in which conventional organic synthesis reactions are performed. This article begins with a brief summary of the most recent advances in the application of enzymes in nonaqueous catalysis. Then a detailed description of synthesis of enzyme nanogel via aqueous two-step in situ polymerization to form an enzyme nanogel is provided. The factors that influence the preparation of enzyme nanogels and increase in enzyme conformational stability are studied by molecular dynamics simulation and multidimensional structural characterization, including transmission electron microscopy, fluorescence emission spectroscopy, and enzymatic catalysis. The potential application of the enzyme nanogel in nonaqueous catalysis at high temperatures was tested using lipase as the model enzyme in reactions conducted in oil–water biphasic media, anhydrous dimethyl sulfoxide (DMSO), and under high vacuum at 95 °C.
2.67.2 Nonaqueous Enzymatic Catalysis Modern biotechnology has generated great interest in the application of biomolecules such as enzymes as catalysts in the production of chemicals, materials, and fuels. The advantages of enzymatic catalysis, including the high efficiency, mild reaction conditions, and environmentally benign reactions, can be exploited by using such biomolecules [1]. Unfortunately, the majority of
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chemical synthesis reactions is presently conducted in the organic phase and at moderate or high temperatures. These conditions differ distinctly from the aqueous physiological environment in which enzymes function stably in their natural ways. For synthetic reactions such as esterification in which water is produced, the nonaqueous phase has been used to prevent the negative inhibition of the reaction by water. On the other hand, most organic species are not soluble in water; therefore, organic solvents have to be used to dissolve such reactants. Klibanov and co-workers observed that when lipases were used to catalyze esterification reactions at 100 °C in anhydrous organic media, their thermal stability was much higher than that in aqueous media [2]. This greatly stimulated research on nonaqueous enzymatic catalysis, which was observed by the increasing amount of literature over the past decades on improved substrate selectivity, regioselectivity, and chemoselectivity of enzymatic catalysis in organic solvents [3]. Increasing efforts have thus been directed toward using an enzyme, a combination of several kinds of enzymes, or an integrated chemical and enzymatic approach to produce fine chemicals, pharmaceuticals, and other complex molecules in which the stereoselectivity of the enzyme plays an essential role. For example, BASF has applied a lipase catalyst for the resolution of racemic alcohol and amine compounds [1]. Nonaqueous enzymatic catalysis can be categorized into four groups based on the composition of the reaction media: (1) water/ water-miscible organic solvent; (2) water/water-immiscible organic solvent; (3) reverse micelles; and (4) neat organic solvent. For reactions carried out in water/water-miscible organic solvents, there is a threshold concentration of the hydrophilic solvent in the aqueous phase, and above this concentration, the enzyme can be seriously deactivated by the organic solvent. For reactions conducted in water/water-immiscible organic solvents, the enzymes partition to the aqueous phase and catalyze the reaction at the interface. Enzymes are generally more stable in neat hydrophobic organic solvents (water-immiscible organic solvents) than in water, while they are very unstable in neat hydrophilic organic solvents (water-miscible organic solvents) such as DMSO or dimethyl formamide (DMF). Although DMSO and DMF are regarded as ‘universal solvents’ for organic synthesis, they are strong denaturants for most enzymes because they dissociate the tertiary structure of the enzyme and strip essential water molecules from the enzyme surface, leading to enzyme unfolding. Klibanov and co-workers established that a native enzyme, a protease from Bacillus subtilis, was active in anhydrous DMF. They predicted unprecedented possibilities for carrying out enzymatic catalysis in organic solvents [4]. To date, very few enzymatic catalysis processes have been realized in DMSO or DMF. Exploration of novel ways to carry out enzymatic reactions in organic solvents or the water–organic solvent media is required, considering the immense potential of enzymatic catalysis.
2.67.3 Enzymes in Nanostructures Studies on enzyme immobilization as a means to enhance enzyme stability and reusability in industrial practice began in the 1960s. In recent years, there have been increasing efforts to develop novel methods for preparing nanostructured enzyme catalysts [5]. These methods include immobilizing enzymes in inorganic or organic nanoparticles, encapsulating enzymes in mesoporous materials, using sol–gel materials, and preparing cross-linked enzyme crystals and cross-linked enzyme aggregates. In comparison with other established effective bulk materials used for enzyme immobilization, nanostructured materials provide additional advantages such as a large surface area, high loading capacity for incorporating molecules, well-tailored microenvironment for maintaining high enzyme activity and stability at high temperatures and in the presence of organic solvents, and low mass transfer resistance within the interior of materials. All of these features are of potential interest in industrial applications, which require stable and efficient catalysts.
2.67.4 Preparation of Enzyme Nanogels 2.67.4.1
Synthetic Procedure
Liu and co-workers established a two-step procedure to encapsulate a protein in a polyacrylamide nanogel [6]. In the first step, vinyl groups were generated on the protein surface mainly by the reaction between N-acryloxysuccinimide (NAS) and an amino group of lysine, as shown in Figure 1. The second step involved aqueous in situ polymerization in which the acryloylated protein was encapsulated. The effectiveness of this method in preparing protein nanogels was tested with horseradish peroxidase (HRP) [6], carbonic anhydrase (CAB) [7], and lipase [8]. In situ polymerization essentially ensures a high yield of the enzyme nanogel, while
NAS
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Enzyme Figure 1 Preparation of the enzyme nanogel.
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Figure 2 TEM image of the lipase (from Candida rugosa ) nanogel.
the aqueous phase provides a friendly environment for the fabrication of enzyme. These conditions lead to high encapsulation yields and enzyme activity. Moreover, the size of the enzyme nanogel can be conveniently tuned by regulating the monomer loading, and enzyme nanogels ranging in size from 10 to 50 nm can be obtained. Figure 2 shows the transmission electron microscopy (TEM) image of a Candida rugosa lipase nanogel prepared by this method. The average diameter of the lipase nanogel was 25 nm. Figure 3 gives the dynamic light-scattering analysis of the lipase nanogel, from which the average diameter of the nanogel is determined as 30 nm.
2.67.4.2 2.67.4.2.1
Optimization of the Synthetic Reactions Used for Lipase Nanogel Preparation The pH of the acryloylation reaction
In principle, an alkaline pH (∼pH 8.0–9.0) enhances the nucleophilicity of the amino group of the protein, which in turn favors its chemical reaction with NAS. However, for a given enzyme, the reaction pH has to be determined by considering the enzyme’s stability and activity. In an earlier study, we observed that a high pH favored the chemical modification of the amino group of lipase but led to unexpectedly low residual activity of the lipase [8]. The optimal pH value for the acryloylation of lipase with NAS was determined to be 4.0, and the residual activity of the modified lipase was 92% [8].
Lipase nanogel Lipase
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Figure 3 DLS analysis of the lipase (from Candida rugosa ) in aqueous solution.
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2.67.4.2.2
Monomers for in situ polymerization
In the second step of the synthesis in which the acryloylated protein is encapsulated, acrylamide is frequently used as the monomer for the in situ polymerization reaction, while N,N′-methylene bisacrylamide is used as the cross-linker. For the catalysis of a high-molecular-weight substrate or for the synthesis of a high-molecular-weight product, N,N′-methylene bisacrylamide is not added as a cross-linker in the polymerization reaction. In this case, the enzyme is encapsulated by the flexible polymer chain. This makes the encapsulated enzyme more accessible to its high molecular substrate than those synthesized using N,N′-methylene bisacrylamide as the cross-linker.
2.67.4.2.3
Monomer concentration in the polymerization reaction
The monomer concentration in the in situ polymerization reaction will affect the size of the protein nanogel and the yield of protein encapsulated in the nanogel. A high monomer concentration increases the encapsulation yield and results in a thicker gel layer. This enhances the conformational and thermal stability of the enzyme. On the other hand, a thicker gel layer results in higher mass transfer resistance during the enzymatic reaction. Thus, a suitable monomer loading has to be determined by balancing the above mentioned effects. In the case of lipase, addition of acrylamide to a final concentration of 50 mg ml−1 in the in situ polymerization reaction resulted in a lipase nanogel of average diameter 20 nm. The encapsulation yield exceeded 90% and the residual hydrolytic activity was 92%.
2.67.5 Molecular Fundamentals of Enzyme Nanogels Our recent efforts have been directed toward establishing molecular fundamentals of enzyme nanogels by performing molecular dynamics simulation at an all-atom level together with multidimensional structural characterization studies. The objective was to obtain molecular insights into enzyme nanogels, understand the mechanisms underlying the enhanced stability of the enzyme and the enzymatic catalysis reaction, and develop a tool for the design and application of enzyme nanogels.
2.67.5.1
Assembly of Monomers around the Enzyme in Aqueous Solution
The assembly of acrylamide around the lipase was simulated using molecular dynamics simulation at an all-atom level [8]. As shown in Figure 4, the lipase was encapsulated within acrylamide aggregates that could form a gel network by subsequent polymerization. The hydrogen bonding calculations indicated that the preferential formation of hydrogen bonds with the lipase was the major driving force for the assembly of acrylamide monomers around the lipase, as shown in Figure 5. The formation of the assembly in aqueous solution was detected by fluorescence resonance energy transfer (FRET), in which the lipase served as the donor and pyrene was the acceptor. FRET occurred from the lipase to pyrene, resulting in an increase in the fluorescence intensity of pyrene at 370–500 nm and a simultaneous reduction in the fluorescence intensity of lipase at 340 nm, as shown in Figure 6. It was observed that the addition of acrylamide monomers led to an increase in the energy transfer efficiency (ET), indicating that pyrene molecules were pushed to the lipase surface as a result of acrylamide assembly. On the other hand, the polarity of the microenvir onment around pyrene decreased once acrylamide was added, indicating that pyrene molecules are incorporated into a more
Enzyme Water
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Figure 4 Assembly of acrylamide molecules around the lipase surface.
Nanostructured Biocatalysts
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Figure 5 Analysis of hydrogen bonding within the lipase–acrylamide assembly. 1, H-bond between lipase and water in pure water solution; 2, H-bond between lipase and water in acrylamide solution; 3, H-bond between lipase and acrylamide in acrylamide solution; and 4, H-bond between lipase with water and acrylamide in acrylamide solution. Cited from Ge J, Lu DN, Wang J, et al. (2008) Molecular fundamentals of enzyme nanogels. Journal of Physical Chemistry B 112: 14319–14324. With permission from American Chemical Society [8].
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Figure 6 Fluorescence resonance energy transfer (FRET) between lipase and pyrene. Solid line, emission spectrum of lipase; dashed line, emission spectrum of lipase and pyrene. Acrylamide concentration in the lipase–pyrene–acrylamide aqueous solution (w/v%): 1–0%, 2–0.5%, 3–1.0%, 4–1.5%, and 5–2.0%.
hydrophobic microenvironment in the presence of acrylamide. In other words, the original water molecules were expelled by the acrylamide molecules that assembled around the lipase.
2.67.5.2
Enzyme Stabilization via the Polymer Network
Dissociation of the hydrophobic core is a major reason for enzyme deactivation at high temperatures. As shown in Figure 7, molecular simulation studies indicated that the presence of the acrylamide network increased the intramolecular hydrogen bonding in the lipase and, thus, contributed to the enhanced thermal stability of the enzyme. The strengthened intramolecular interactions allowed the encapsulated lipase to withstand higher temperatures, particularly after polymerization that resulted in multipoint linkages with the porous acrylamide network. Moreover, calculations of the radial distribution function (RDF) of water and organic solvents such as DMSO indicated that the presence of the acrylamide network pushed the solvent molecules away from the surface of the lipase, resulting in enhanced stability of the lipase in the organic solvent, as shown in Figure 8. Finally, an increase in the acrylamide concentration further reduced the root mean square deviation (RMSD) in the presence of DMSO. These results indicated that the gel thickness can be tuned to yield an enzyme nanogel of required stability (Figure 9).
2.67.5.3
Experimental Validation of Enhanced Enzyme Stability
The enhanced stability of the encapsulated enzyme at high temperatures or in the presence of organic solvents was validated using HRP [6] and CAB [7]. The stability of the lipase was studied at high temperatures and in the presence of methanol, as shown in Figure 10.
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Figure 7 Intramolecular hydrogen bonding within the lipase in the presence of acrylamide. Reproduced from Ge J, Lu DN, Wang J, et al. (2008) Molecular fundamentals of enzyme nanogels. Journal of Physical Chemistry B 112: 14319–14324. With permission from American Chemical Society [8].
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Lipase (1) Lipase in 1 nm nanogel (2) Lipase in 3 nm nanogel (3)
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Figure 8 The RDF of DMSO in a lipase–acrylamide aqueous solution. Reproduced from Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. With permission from American Chemical Society [10].
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Figure 9 The RMSD of the lipase in DMSO in the presence of acrylamide. Reproduced from Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. With permission from American Chemical Society [10].
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Figure 10 Stability of the lipase nanogel in methanol.
As shown in Figure 10, the native lipase was almost completely deactivated within half an hour, while the lipase nanogel preserved most of its activity in anhydrous methanol for 3 h. The enhanced tolerance to the hydrophilic solvent can be explained on the basis of molecular simulation studies and was found to be mainly due to the presence of the hydrophilic polyacrylamide network around the lipase surface.
2.67.6 Potential Applications of Enzyme Nanogels as Biocatalysts Lipase was chosen as a model enzyme to examine the potential of enzyme nanogels as robust catalysts for chemical synthesis under harsh conditions. The lipase nanogel was applied to produce (1) biodiesel in methanol–water biphasic media; (2) a dextran-based biodegradable surfactant in anhydrous DMSO; and (3) polyester in the absence of solvent. It was established that the presence of methanol resulted in denaturation of the lipase due to the stripping of essential water. DMSO is known to be denaturing to most enzymes including lipase [9]. Moreover, few enzymes are reported to be active at 95 °C, which is the temperature at which the polycondensation of succinic acid and 1,4-butanediol occurs.
2.67.6.1
Lipase Nanogel-Catalyzed Synthesis of Biodiesel
Enzymatic production of biodiesel (fatty acid methyl esters) by methanolysis of triglycerides using lipase as a catalyst was established as a green process but suffered from high operational costs, which mainly arose from enzyme denaturation by shortchain alcohols such as methanol. In the present study, a genetically modified Aspergillus niger lipase (NS81006 from Novozyme) was encapsulated in a polyacrylamide nanogel of average diameter 25 nm using an aqueous two-step in situ polymerization reaction. The residual esterase and lipase activities were 82 and 68%, respectively, after encapsulation. In comparison to its native counter part, that is, free lipase NS81006, the lipase nanogel showed an operation time that was extended by 2.5-fold at 55 °C and a significantly higher tolerance to methanol. The significantly enhanced stability at high temperature and in the presence of methanol indicates that the lipase nanogel is a promising catalyst for the biphasic enzymatic production of biodiesel.
2.67.6.2
Lipase Nanogel-Catalyzed Synthesis of a Dextran-Based Surfactant
Enzyme-catalyzed synthesis of biocompatible and biodegradable surfactants based on carbohydrates is attractive because it provides a regioselective catalyst that enables a precise molecular structure for the product and simplifies the purification steps for the downstream process. Anhydrous DMSO and DMF were used to dissolve the hydrophilic polysaccharide and hydrophobic components. However, both of these solvents are strong enzyme denaturants. We synthesized the lipase nanogel and used it to catalyze the transesterification reaction between dextran and vinyl decanoate in anhydrous DMSO. The resulting product was dextran–decanoate, a biodegradable surfactant [10]. As shown in Figure 11, the lipase nanogel behaved as a stable catalyst in anhydrous DMSO at 60 °C for 10 days and showed an overwhelming regioselectivity toward the C-2 hydroxyl group in the glucopyranosyl unit of dextran. The degree of substitution was 23%, which was obtained using the lipase nanogel as the catalyst. By contrast, the use of unmodified lipase resulted in a degree of substitution less than 3%.
2.67.6.3
Lipase Nanogel-Catalyzed Synthesis of Polyester
In comparison with the chemically catalyzed synthesis of polyesters, enzyme-catalyzed reactions may provide an environmentally friendly method for producing biodegradable and biocompatible polyesters, which may be applied for medical purposes [11]. Moreover, lipase-catalyzed polymerization is a straightforward method for synthesizing linear polyesters with pendant functional
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Figure 11 Lipase nanogel-catalyzed synthesis of a dextran-based surfactant. Conversion curves of transesterification between dextran and VD catalyzed by lipase and lipase nanogel: (1) native lipase, 60 °C; (2) lipase nanogel, 60 °C; and (3) mixture of lipase and polyacrylamide, 60 °C. Reproduced from Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. With permission from American Chemical Society [10].
groups. Succinic acid and 1,4-butanediol are convenient substrates for the synthesis of poly(butylene succinate) (PBS), an important kind of polyester. When the C. rugosa lipase nanogel of average diameter 20 nm was used as the catalyst, the poly condensation of succinic acid and 1,4-butanediol was performed at 95 °C under high vacuum (<10 mm Hg). After carrying out the reaction for 72 h under these conditions, biodegradable PBS with an Mn of 1820 g mol−1, Mw of 1900 g mol−1, and melting point of 107.2 °C was obtained. These results showed that the lipase nanogel could be used for the non-solvent-based synthesis of polyesters.
2.67.7 Summary In this article, we have summarized the latest advances in both nonaqueous enzymatic synthesis and nanostructured enzyme catalysts that exhibit high stability while retaining their original catalytic activity. An aqueous two-step in situ polymerization method for encapsulating enzyme nanogel has been detailed, which produced a robust enzyme catalyst that can function under adverse conditions such as high temperatures and the presence of organic solvents. The molecular fundamentals of the enzyme nanogel were established using molecular dynamics simulations at an all-atom level together with multidimensional structural characterization studies. It was shown that hydrogen bonding with the protein drives the monomer to form assemblies around the protein, which can then polymerize into enzyme nanogels via subsequent in situ polymerization. The polymer network not only enhances the thermal stability of the enzyme but also inhibits the stripping of essential water from the enzyme surface by the polar solvent. This gives the encapsulated enzyme significantly enhanced stability at high temperatures and in the presence of organic solvents. The reactions performed in oil–water biphasic media, anhydrous DMSO, and nonsolvent media using lipase nanogels as the catalyst illustrate the immense potential of enzyme nanogels as robust catalysts for chemical synthesis.
References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11]
Schmid A, Dordick JS, Hauer B, et al. (2001) Industrial biocatalysis today and tomorrow. Nature 409: 258–268. Zaks A and Klibanov AM (1984) Enzymatic catalysis in organic media at 100 °C. Science 224: 1249–1251. Klibanov AM (2001) Improving enzymes by using them in organic solvents. Nature 409: 241–246. Riva S, Chopineau J, Kieboom APG, and Klibanov AM (1988) Protease-catalyzed regioselective esterification of sugars and related compounds in anhydrous dimethylformamide. Journal of the American Chemical Society 110: 584–589. Kim JB, Grate JW, and Wang P (2008) Nanobiocatalysis and its potential applications. Trends in Biotechnology 26: 639–646. Yan M, Ge J, Liu Z, and Ouyang PK (2006) Encapsulation of single enzyme in nanogel with enhanced biocatalytic activity and stability. Journal of the American Chemical Society 128: 11008–11009. Yan M, Liu ZX, Lu DN, and Liu Z (2007) Fabrication of single carbonic anhydrase nanogel against denaturation and aggregation at high temperature. Biomacromolecules 8: 560–565. Ge J, Lu DN, Wang J, et al. (2008) Molecular fundamentals of enzyme nanogels. Journal of Physical Chemistry B 112: 14319–14324. Zaks A and Klibanov AM (1985) Enzyme-catalyzed processes in organic solvents. Proceedings of the National Academy of Sciences of the United States of America 82: 3192–3196. Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. Kobayashi S (2009) Recent developments in lipase-catalyzed synthesis of polyesters. Macromolecular Rapid Communications 30: 237–266.
Nanostructured Biocatalysts
J Ge, D Lu, M Yan, and Z Liu, Tsinghua University, Beijing, China © 2011 Elsevier B.V. All rights reserved.
2.67.1 2.67.2 2.67.3 2.67.4 2.67.4.1 2.67.4.2 2.67.4.2.1 2.67.4.2.2 2.67.4.2.3 2.67.5 2.67.5.1 2.67.5.2 2.67.5.3 2.67.6 2.67.6.1 2.67.6.2 2.67.6.3 2.67.7 References
Introduction Nonaqueous Enzymatic Catalysis Enzymes in Nanostructures Preparation of Enzyme Nanogels Synthetic Procedure Optimization of the Synthetic Reactions Used for Lipase Nanogel Preparation The pH of the acryloylation reaction Monomers for in situ polymerization Monomer concentration in the polymerization reaction Molecular Fundamentals of Enzyme Nanogels Assembly of Monomers around the Enzyme in Aqueous Solution Enzyme Stabilization via the Polymer Network Experimental Validation of Enhanced Enzyme Stability Potential Applications of Enzyme Nanogels as Biocatalysts Lipase Nanogel-Catalyzed Synthesis of Biodiesel Lipase Nanogel-Catalyzed Synthesis of a Dextran-Based Surfactant Lipase Nanogel-Catalyzed Synthesis of Polyester Summary
Glossary enzymatic catalysis The chemical reactions catalyzed by an enzyme that naturally catalyzes reactions in a biological system. enzyme stabilization The physical or chemical methods to enhance enzyme stability under adverse conditions such as high temperature or the presence of denaturing solvents. enzyme modification The chemical methods that introduce additional structure to an enzyme or to enhance enzyme stability and/or activity.
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enzyme nanogel The transparent polymer matrix with diameter below 100 nm, which is obtained by in situ polymerization from the enzyme surface. nanostructured enzyme catalyst The materials below 100 nm in dimension(s) such as nanobeads and nanofibers, which display the activity of the incorporated enzyme.
2.67.1 Introduction Enzyme stabilization is of fundamental importance for extending the scope of enzymatic catalysis to nonaqueous media in which conventional organic synthesis reactions are performed. This article begins with a brief summary of the most recent advances in the application of enzymes in nonaqueous catalysis. Then a detailed description of synthesis of enzyme nanogel via aqueous two-step in situ polymerization to form an enzyme nanogel is provided. The factors that influence the preparation of enzyme nanogels and increase in enzyme conformational stability are studied by molecular dynamics simulation and multidimensional structural characterization, including transmission electron microscopy, fluorescence emission spectroscopy, and enzymatic catalysis. The potential application of the enzyme nanogel in nonaqueous catalysis at high temperatures was tested using lipase as the model enzyme in reactions conducted in oil–water biphasic media, anhydrous dimethyl sulfoxide (DMSO), and under high vacuum at 95 °C.
2.67.2 Nonaqueous Enzymatic Catalysis Modern biotechnology has generated great interest in the application of biomolecules such as enzymes as catalysts in the production of chemicals, materials, and fuels. The advantages of enzymatic catalysis, including the high efficiency, mild reaction conditions, and environmentally benign reactions, can be exploited by using such biomolecules [1]. Unfortunately, the majority of
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chemical synthesis reactions is presently conducted in the organic phase and at moderate or high temperatures. These conditions differ distinctly from the aqueous physiological environment in which enzymes function stably in their natural ways. For synthetic reactions such as esterification in which water is produced, the nonaqueous phase has been used to prevent the negative inhibition of the reaction by water. On the other hand, most organic species are not soluble in water; therefore, organic solvents have to be used to dissolve such reactants. Klibanov and co-workers observed that when lipases were used to catalyze esterification reactions at 100 °C in anhydrous organic media, their thermal stability was much higher than that in aqueous media [2]. This greatly stimulated research on nonaqueous enzymatic catalysis, which was observed by the increasing amount of literature over the past decades on improved substrate selectivity, regioselectivity, and chemoselectivity of enzymatic catalysis in organic solvents [3]. Increasing efforts have thus been directed toward using an enzyme, a combination of several kinds of enzymes, or an integrated chemical and enzymatic approach to produce fine chemicals, pharmaceuticals, and other complex molecules in which the stereoselectivity of the enzyme plays an essential role. For example, BASF has applied a lipase catalyst for the resolution of racemic alcohol and amine compounds [1]. Nonaqueous enzymatic catalysis can be categorized into four groups based on the composition of the reaction media: (1) water/ water-miscible organic solvent; (2) water/water-immiscible organic solvent; (3) reverse micelles; and (4) neat organic solvent. For reactions carried out in water/water-miscible organic solvents, there is a threshold concentration of the hydrophilic solvent in the aqueous phase, and above this concentration, the enzyme can be seriously deactivated by the organic solvent. For reactions conducted in water/water-immiscible organic solvents, the enzymes partition to the aqueous phase and catalyze the reaction at the interface. Enzymes are generally more stable in neat hydrophobic organic solvents (water-immiscible organic solvents) than in water, while they are very unstable in neat hydrophilic organic solvents (water-miscible organic solvents) such as DMSO or dimethyl formamide (DMF). Although DMSO and DMF are regarded as ‘universal solvents’ for organic synthesis, they are strong denaturants for most enzymes because they dissociate the tertiary structure of the enzyme and strip essential water molecules from the enzyme surface, leading to enzyme unfolding. Klibanov and co-workers established that a native enzyme, a protease from Bacillus subtilis, was active in anhydrous DMF. They predicted unprecedented possibilities for carrying out enzymatic catalysis in organic solvents [4]. To date, very few enzymatic catalysis processes have been realized in DMSO or DMF. Exploration of novel ways to carry out enzymatic reactions in organic solvents or the water–organic solvent media is required, considering the immense potential of enzymatic catalysis.
2.67.3 Enzymes in Nanostructures Studies on enzyme immobilization as a means to enhance enzyme stability and reusability in industrial practice began in the 1960s. In recent years, there have been increasing efforts to develop novel methods for preparing nanostructured enzyme catalysts [5]. These methods include immobilizing enzymes in inorganic or organic nanoparticles, encapsulating enzymes in mesoporous materials, using sol–gel materials, and preparing cross-linked enzyme crystals and cross-linked enzyme aggregates. In comparison with other established effective bulk materials used for enzyme immobilization, nanostructured materials provide additional advantages such as a large surface area, high loading capacity for incorporating molecules, well-tailored microenvironment for maintaining high enzyme activity and stability at high temperatures and in the presence of organic solvents, and low mass transfer resistance within the interior of materials. All of these features are of potential interest in industrial applications, which require stable and efficient catalysts.
2.67.4 Preparation of Enzyme Nanogels 2.67.4.1
Synthetic Procedure
Liu and co-workers established a two-step procedure to encapsulate a protein in a polyacrylamide nanogel [6]. In the first step, vinyl groups were generated on the protein surface mainly by the reaction between N-acryloxysuccinimide (NAS) and an amino group of lysine, as shown in Figure 1. The second step involved aqueous in situ polymerization in which the acryloylated protein was encapsulated. The effectiveness of this method in preparing protein nanogels was tested with horseradish peroxidase (HRP) [6], carbonic anhydrase (CAB) [7], and lipase [8]. In situ polymerization essentially ensures a high yield of the enzyme nanogel, while
NAS
Monomer Polymerization
Enzyme Figure 1 Preparation of the enzyme nanogel.
Enzyme nanogel
Nanostructured Biocatalysts
EN4657 X50 K 120.0 KV
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Figure 2 TEM image of the lipase (from Candida rugosa ) nanogel.
the aqueous phase provides a friendly environment for the fabrication of enzyme. These conditions lead to high encapsulation yields and enzyme activity. Moreover, the size of the enzyme nanogel can be conveniently tuned by regulating the monomer loading, and enzyme nanogels ranging in size from 10 to 50 nm can be obtained. Figure 2 shows the transmission electron microscopy (TEM) image of a Candida rugosa lipase nanogel prepared by this method. The average diameter of the lipase nanogel was 25 nm. Figure 3 gives the dynamic light-scattering analysis of the lipase nanogel, from which the average diameter of the nanogel is determined as 30 nm.
2.67.4.2 2.67.4.2.1
Optimization of the Synthetic Reactions Used for Lipase Nanogel Preparation The pH of the acryloylation reaction
In principle, an alkaline pH (∼pH 8.0–9.0) enhances the nucleophilicity of the amino group of the protein, which in turn favors its chemical reaction with NAS. However, for a given enzyme, the reaction pH has to be determined by considering the enzyme’s stability and activity. In an earlier study, we observed that a high pH favored the chemical modification of the amino group of lipase but led to unexpectedly low residual activity of the lipase [8]. The optimal pH value for the acryloylation of lipase with NAS was determined to be 4.0, and the residual activity of the modified lipase was 92% [8].
Lipase nanogel Lipase
1
10 Rh (nm)
Figure 3 DLS analysis of the lipase (from Candida rugosa ) in aqueous solution.
100
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Other Considerations
2.67.4.2.2
Monomers for in situ polymerization
In the second step of the synthesis in which the acryloylated protein is encapsulated, acrylamide is frequently used as the monomer for the in situ polymerization reaction, while N,N′-methylene bisacrylamide is used as the cross-linker. For the catalysis of a high-molecular-weight substrate or for the synthesis of a high-molecular-weight product, N,N′-methylene bisacrylamide is not added as a cross-linker in the polymerization reaction. In this case, the enzyme is encapsulated by the flexible polymer chain. This makes the encapsulated enzyme more accessible to its high molecular substrate than those synthesized using N,N′-methylene bisacrylamide as the cross-linker.
2.67.4.2.3
Monomer concentration in the polymerization reaction
The monomer concentration in the in situ polymerization reaction will affect the size of the protein nanogel and the yield of protein encapsulated in the nanogel. A high monomer concentration increases the encapsulation yield and results in a thicker gel layer. This enhances the conformational and thermal stability of the enzyme. On the other hand, a thicker gel layer results in higher mass transfer resistance during the enzymatic reaction. Thus, a suitable monomer loading has to be determined by balancing the above mentioned effects. In the case of lipase, addition of acrylamide to a final concentration of 50 mg ml−1 in the in situ polymerization reaction resulted in a lipase nanogel of average diameter 20 nm. The encapsulation yield exceeded 90% and the residual hydrolytic activity was 92%.
2.67.5 Molecular Fundamentals of Enzyme Nanogels Our recent efforts have been directed toward establishing molecular fundamentals of enzyme nanogels by performing molecular dynamics simulation at an all-atom level together with multidimensional structural characterization studies. The objective was to obtain molecular insights into enzyme nanogels, understand the mechanisms underlying the enhanced stability of the enzyme and the enzymatic catalysis reaction, and develop a tool for the design and application of enzyme nanogels.
2.67.5.1
Assembly of Monomers around the Enzyme in Aqueous Solution
The assembly of acrylamide around the lipase was simulated using molecular dynamics simulation at an all-atom level [8]. As shown in Figure 4, the lipase was encapsulated within acrylamide aggregates that could form a gel network by subsequent polymerization. The hydrogen bonding calculations indicated that the preferential formation of hydrogen bonds with the lipase was the major driving force for the assembly of acrylamide monomers around the lipase, as shown in Figure 5. The formation of the assembly in aqueous solution was detected by fluorescence resonance energy transfer (FRET), in which the lipase served as the donor and pyrene was the acceptor. FRET occurred from the lipase to pyrene, resulting in an increase in the fluorescence intensity of pyrene at 370–500 nm and a simultaneous reduction in the fluorescence intensity of lipase at 340 nm, as shown in Figure 6. It was observed that the addition of acrylamide monomers led to an increase in the energy transfer efficiency (ET), indicating that pyrene molecules were pushed to the lipase surface as a result of acrylamide assembly. On the other hand, the polarity of the microenvir onment around pyrene decreased once acrylamide was added, indicating that pyrene molecules are incorporated into a more
Enzyme Water
Monomer
Figure 4 Assembly of acrylamide molecules around the lipase surface.
Nanostructured Biocatalysts
929
800
H-bond
600
400
200
0
1
2
3
4
Figure 5 Analysis of hydrogen bonding within the lipase–acrylamide assembly. 1, H-bond between lipase and water in pure water solution; 2, H-bond between lipase and water in acrylamide solution; 3, H-bond between lipase and acrylamide in acrylamide solution; and 4, H-bond between lipase with water and acrylamide in acrylamide solution. Cited from Ge J, Lu DN, Wang J, et al. (2008) Molecular fundamentals of enzyme nanogels. Journal of Physical Chemistry B 112: 14319–14324. With permission from American Chemical Society [8].
1200
Intensity
1000 800
1
600
2
400
3 4
200 5 0 300
350
400 450 Wavelength (nm)
500
550
Figure 6 Fluorescence resonance energy transfer (FRET) between lipase and pyrene. Solid line, emission spectrum of lipase; dashed line, emission spectrum of lipase and pyrene. Acrylamide concentration in the lipase–pyrene–acrylamide aqueous solution (w/v%): 1–0%, 2–0.5%, 3–1.0%, 4–1.5%, and 5–2.0%.
hydrophobic microenvironment in the presence of acrylamide. In other words, the original water molecules were expelled by the acrylamide molecules that assembled around the lipase.
2.67.5.2
Enzyme Stabilization via the Polymer Network
Dissociation of the hydrophobic core is a major reason for enzyme deactivation at high temperatures. As shown in Figure 7, molecular simulation studies indicated that the presence of the acrylamide network increased the intramolecular hydrogen bonding in the lipase and, thus, contributed to the enhanced thermal stability of the enzyme. The strengthened intramolecular interactions allowed the encapsulated lipase to withstand higher temperatures, particularly after polymerization that resulted in multipoint linkages with the porous acrylamide network. Moreover, calculations of the radial distribution function (RDF) of water and organic solvents such as DMSO indicated that the presence of the acrylamide network pushed the solvent molecules away from the surface of the lipase, resulting in enhanced stability of the lipase in the organic solvent, as shown in Figure 8. Finally, an increase in the acrylamide concentration further reduced the root mean square deviation (RMSD) in the presence of DMSO. These results indicated that the gel thickness can be tuned to yield an enzyme nanogel of required stability (Figure 9).
2.67.5.3
Experimental Validation of Enhanced Enzyme Stability
The enhanced stability of the encapsulated enzyme at high temperatures or in the presence of organic solvents was validated using HRP [6] and CAB [7]. The stability of the lipase was studied at high temperatures and in the presence of methanol, as shown in Figure 10.
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Other Considerations
440 Lipase nanogel Intramolecular H-bond
420 400 380 360 340 Lipase 320
0
200
400 600 Time (ps)
800
1000
Figure 7 Intramolecular hydrogen bonding within the lipase in the presence of acrylamide. Reproduced from Ge J, Lu DN, Wang J, et al. (2008) Molecular fundamentals of enzyme nanogels. Journal of Physical Chemistry B 112: 14319–14324. With permission from American Chemical Society [8].
1.2
Lipase (1) Lipase in 1 nm nanogel (2) Lipase in 3 nm nanogel (3)
1.0
gAB(r)
.8 3
.6
2
1
.4
.2
0.0
0
2
4 r (nm)
6
Figure 8 The RDF of DMSO in a lipase–acrylamide aqueous solution. Reproduced from Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. With permission from American Chemical Society [10].
1.2 1.0
RMSD
.8 .6
Lipase In 1 nm gel In 3 nm gel
.4
.2
0.0
0
1000
2000 Time (ps)
3000
Figure 9 The RMSD of the lipase in DMSO in the presence of acrylamide. Reproduced from Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. With permission from American Chemical Society [10].
Nanostructured Biocatalysts
931
140 Lipase Relative activity (%)
120
Lipase nanogel
100 80 60 40 20 0 0
40
80 120 Time (min)
160
200
Figure 10 Stability of the lipase nanogel in methanol.
As shown in Figure 10, the native lipase was almost completely deactivated within half an hour, while the lipase nanogel preserved most of its activity in anhydrous methanol for 3 h. The enhanced tolerance to the hydrophilic solvent can be explained on the basis of molecular simulation studies and was found to be mainly due to the presence of the hydrophilic polyacrylamide network around the lipase surface.
2.67.6 Potential Applications of Enzyme Nanogels as Biocatalysts Lipase was chosen as a model enzyme to examine the potential of enzyme nanogels as robust catalysts for chemical synthesis under harsh conditions. The lipase nanogel was applied to produce (1) biodiesel in methanol–water biphasic media; (2) a dextran-based biodegradable surfactant in anhydrous DMSO; and (3) polyester in the absence of solvent. It was established that the presence of methanol resulted in denaturation of the lipase due to the stripping of essential water. DMSO is known to be denaturing to most enzymes including lipase [9]. Moreover, few enzymes are reported to be active at 95 °C, which is the temperature at which the polycondensation of succinic acid and 1,4-butanediol occurs.
2.67.6.1
Lipase Nanogel-Catalyzed Synthesis of Biodiesel
Enzymatic production of biodiesel (fatty acid methyl esters) by methanolysis of triglycerides using lipase as a catalyst was established as a green process but suffered from high operational costs, which mainly arose from enzyme denaturation by shortchain alcohols such as methanol. In the present study, a genetically modified Aspergillus niger lipase (NS81006 from Novozyme) was encapsulated in a polyacrylamide nanogel of average diameter 25 nm using an aqueous two-step in situ polymerization reaction. The residual esterase and lipase activities were 82 and 68%, respectively, after encapsulation. In comparison to its native counter part, that is, free lipase NS81006, the lipase nanogel showed an operation time that was extended by 2.5-fold at 55 °C and a significantly higher tolerance to methanol. The significantly enhanced stability at high temperature and in the presence of methanol indicates that the lipase nanogel is a promising catalyst for the biphasic enzymatic production of biodiesel.
2.67.6.2
Lipase Nanogel-Catalyzed Synthesis of a Dextran-Based Surfactant
Enzyme-catalyzed synthesis of biocompatible and biodegradable surfactants based on carbohydrates is attractive because it provides a regioselective catalyst that enables a precise molecular structure for the product and simplifies the purification steps for the downstream process. Anhydrous DMSO and DMF were used to dissolve the hydrophilic polysaccharide and hydrophobic components. However, both of these solvents are strong enzyme denaturants. We synthesized the lipase nanogel and used it to catalyze the transesterification reaction between dextran and vinyl decanoate in anhydrous DMSO. The resulting product was dextran–decanoate, a biodegradable surfactant [10]. As shown in Figure 11, the lipase nanogel behaved as a stable catalyst in anhydrous DMSO at 60 °C for 10 days and showed an overwhelming regioselectivity toward the C-2 hydroxyl group in the glucopyranosyl unit of dextran. The degree of substitution was 23%, which was obtained using the lipase nanogel as the catalyst. By contrast, the use of unmodified lipase resulted in a degree of substitution less than 3%.
2.67.6.3
Lipase Nanogel-Catalyzed Synthesis of Polyester
In comparison with the chemically catalyzed synthesis of polyesters, enzyme-catalyzed reactions may provide an environmentally friendly method for producing biodegradable and biocompatible polyesters, which may be applied for medical purposes [11]. Moreover, lipase-catalyzed polymerization is a straightforward method for synthesizing linear polyesters with pendant functional
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Other Considerations
Substitution degree (%)
30 25
(1)
20 15 10 5
(2) (3)
0 0
50
100
150 Time (h)
200
250
Figure 11 Lipase nanogel-catalyzed synthesis of a dextran-based surfactant. Conversion curves of transesterification between dextran and VD catalyzed by lipase and lipase nanogel: (1) native lipase, 60 °C; (2) lipase nanogel, 60 °C; and (3) mixture of lipase and polyacrylamide, 60 °C. Reproduced from Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. With permission from American Chemical Society [10].
groups. Succinic acid and 1,4-butanediol are convenient substrates for the synthesis of poly(butylene succinate) (PBS), an important kind of polyester. When the C. rugosa lipase nanogel of average diameter 20 nm was used as the catalyst, the poly condensation of succinic acid and 1,4-butanediol was performed at 95 °C under high vacuum (<10 mm Hg). After carrying out the reaction for 72 h under these conditions, biodegradable PBS with an Mn of 1820 g mol−1, Mw of 1900 g mol−1, and melting point of 107.2 °C was obtained. These results showed that the lipase nanogel could be used for the non-solvent-based synthesis of polyesters.
2.67.7 Summary In this article, we have summarized the latest advances in both nonaqueous enzymatic synthesis and nanostructured enzyme catalysts that exhibit high stability while retaining their original catalytic activity. An aqueous two-step in situ polymerization method for encapsulating enzyme nanogel has been detailed, which produced a robust enzyme catalyst that can function under adverse conditions such as high temperatures and the presence of organic solvents. The molecular fundamentals of the enzyme nanogel were established using molecular dynamics simulations at an all-atom level together with multidimensional structural characterization studies. It was shown that hydrogen bonding with the protein drives the monomer to form assemblies around the protein, which can then polymerize into enzyme nanogels via subsequent in situ polymerization. The polymer network not only enhances the thermal stability of the enzyme but also inhibits the stripping of essential water from the enzyme surface by the polar solvent. This gives the encapsulated enzyme significantly enhanced stability at high temperatures and in the presence of organic solvents. The reactions performed in oil–water biphasic media, anhydrous DMSO, and nonsolvent media using lipase nanogels as the catalyst illustrate the immense potential of enzyme nanogels as robust catalysts for chemical synthesis.
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Schmid A, Dordick JS, Hauer B, et al. (2001) Industrial biocatalysis today and tomorrow. Nature 409: 258–268. Zaks A and Klibanov AM (1984) Enzymatic catalysis in organic media at 100 °C. Science 224: 1249–1251. Klibanov AM (2001) Improving enzymes by using them in organic solvents. Nature 409: 241–246. Riva S, Chopineau J, Kieboom APG, and Klibanov AM (1988) Protease-catalyzed regioselective esterification of sugars and related compounds in anhydrous dimethylformamide. Journal of the American Chemical Society 110: 584–589. Kim JB, Grate JW, and Wang P (2008) Nanobiocatalysis and its potential applications. Trends in Biotechnology 26: 639–646. Yan M, Ge J, Liu Z, and Ouyang PK (2006) Encapsulation of single enzyme in nanogel with enhanced biocatalytic activity and stability. Journal of the American Chemical Society 128: 11008–11009. Yan M, Liu ZX, Lu DN, and Liu Z (2007) Fabrication of single carbonic anhydrase nanogel against denaturation and aggregation at high temperature. Biomacromolecules 8: 560–565. Ge J, Lu DN, Wang J, et al. (2008) Molecular fundamentals of enzyme nanogels. Journal of Physical Chemistry B 112: 14319–14324. Zaks A and Klibanov AM (1985) Enzyme-catalyzed processes in organic solvents. Proceedings of the National Academy of Sciences of the United States of America 82: 3192–3196. Ge J, Lu DN, Wang J, and Liu Z (2009) Lipase nanogel catalyzed transesterification in anhydrous dimethyl sulfoxide. Biomacromolecules 10: 1612–1618. Kobayashi S (2009) Recent developments in lipase-catalyzed synthesis of polyesters. Macromolecular Rapid Communications 30: 237–266.