NANO LETTERS
Amphiphilic Network as Nanoreactor for Enzymes in Organic Solvents
2005 Vol. 5, No. 1 45-48
Nico Bruns and Joerg C. Tiller* Freiburg Materials Research Center (FMF) and Institute of Macromolecular Chemistry, Albert-Ludwigs-UniVersity Freiburg, Stefan-Meier-Strasse 21, 79104 Freiburg, Germany Received September 27, 2004; Revised Manuscript Received November 12, 2004
ABSTRACT Enzymes are powerful biocatalysts that work naturally in water but are also active in organic solvents. Here, we present a nanophaseseparated amphiphilic network, where an enzyme is entrapped into its hydrophilic domains. A substrate that diffuses into the other, hydrophobic, phase of such a network can access the biocatalyst via the extremely large interface. Entrapped horseradish peroxidase and chloroperoxidase showed dramatically increased activity and operational stability compared to the native enzymes.
Enzymes might become the catalysts of the future in organic synthesis. Their enormous potential as stereoselective catalysts in asymmetric synthesis is increasingly exploited1 due to the ever-growing demand in enantiopure pharmaceuticals.2 The technological and synthetic utility of these biocatalysts can be enhanced greatly by using them in organic solvents rather than in their natural aqueous reaction media.3-5 However, and although the environmental advantages of enzymes over metal-organic catalysts are obvious, only few reactions in organic media are catalyzed by these biomolecules on an industrial scale, e.g., racemic resolution of alcohols and amines by lipases.6 Most often a commercial application fails because of the low activity and an often decreased stability of enzymes in organic media. Low catalytic activities are mainly due to the insolubility of most enzymes in organic solvents.7 The biocatalysts are usually used in suspension in these media, resulting in a heterogeneous catalysis. The enhancement of enzyme activity in organic media is of great current interest.8 Besides the most elaborate proteinengineering methods, the activity of biocatalysts can be enhanced by additives. Examples are poly(ethylene glycol) (PEG)9 and lipids,10 acting as structure-preserving lyoprotectants and solubilizing agents, respectively. Unfortunately, additives represent impurities that are difficult to remove from the products. Another way to increase enzyme activity is to modify the biocatalyst by either grafting a polymer, e.g. PEG,11 onto it or covalently binding it into a polymer network.12,13 Both ways require a rather elaborate modification of enzymes. The use of macro- or microporous supports, e.g. silica, glass or polymer beads,14 as carriers for enzymes is another widely used approach, but the amount of accessible enzyme molecules is limited by the surface area of the * Corresponding author. Fax: (+49) 761-2034709. E-mail:joerg.tiller@ fmf.uni-freiburg.de. 10.1021/nl048413b CCC: $30.25 Published on Web 12/04/2004
© 2005 American Chemical Society
Scheme 1. Schematic Depiction of a Nanophase-Separated Amphiphilic Network as Mediative Matrix for Enzyme Activity in Organic Solvents: Loading of Enzyme into the Water-Swollen Hydrophilic Phase and Substrate Conversion in Organic Solvent
particles, as is the case for hydrogel-encapsulated enzymes.15 Finally, encapsulation of biocatalysts in porous sol-gels is a generic approach for enzyme immobilization, being able to enhance catalytic performance of enzymes in organic media.16,17 In all sol-gel-enzyme materials the biocatalyst is accessed by solvents and substrates through the same pores in which it is entrapped. Therefore, the enzyme might be deactivated by the solvent or substrates in high excess. Here we present a technology that provides advantages over these enzyme activation methods. It is based on a nonporous, nanostructured amphiphilic network with an extremely large interface between the networks hydrophilic and hydrophobic phases due to nanophase separation and its peculiar swelling properties.18-20 The network can be easily loaded with an enzyme by simply immersing it into an aqueous solution of the biocatalyst. Therefore, the method does not require additives or elaborate protein modifications. The principle of the network as mediative matrix of enzyme activity is based on the fact that the enzyme can diffuse from its solution into the swollen hydrophilic phase of the network
Figure 1. Synthesis and characterization of amphiphilic networks. (a) Synthesis scheme of PHEA-l-PDMS networks via photopolymerization and subsequent deprotection. (b) AFM phase mode images of PHEA-l-PDMS film-surface and (c) of its cross section, created by a fracture of a film in liquid nitrogen. Hydrophilic, hard PHEA shows light, hydrophobic, soft PDMS shows dark. (d) AFM phase mode image of PHEA-co-PMAA-l-EGDMA (film surface). PHEA shows light, poly(methylacrylate) dark.
(Scheme 1). Upon drying, the phase shrinks, encapsulating the protein into an enzyme-friendly environment.21 The hydrophobic phase swells in a nonpolar organic solvent, allowing a dissolved substrate to enter the network and getting in contact with the enzyme through the interface between the hydrophilic and hydrophobic phase. The formed product can diffuse out of the network. We chose a nanophase-separated amphiphilic network on the basis of poly(2-hydroxyethyl acrylate) (PHEA) and bifunctional poly(dimethylsiloxane) (PDMS) macromonomers [R,ω-methacryloxypropyl poly(dimethyl siloxane) (Mn ) 5200 g mol-1)], consisting of 77 wt % PHEA and 23 wt % PDMS (referred as PHEA-l-PDMS 1) for our experiments. It was synthesized by UV-initiated radical copolymerization as a polymer film of some 20 µm in thickness, covalently attached to glass substrates. As seen in Figures 1b and 1c, the network shows phase separation in the bulk as well as on the surface of the polymeric film. The latter is the basis of the success of the network, because free access to its interior for enzymes (in water) as well as substrates (in organic solvent) is essential for the function of the network. The AFM pictures indicate a sponge-like morphology where PDMS phases are enclosed in a PHEA phase which forms the walls of the sponge structure. On the surface and in bulk the hydrophobic domains are between 12 and 22 nm in 46
diameter. The hydrophilic walls have a thickness of 2 to 7 nm on the surface, whereas they form thicker structures in the bulk of the film. There the thickness ranges between 5 and 20 nm. For the proposed application, the network domain sizes must be carefully considered. The size of the hydrophilic domains should be at least as big as the enzyme (i.e., several nanometers) to allow a good encapsulation and protection of the enzyme. On the other hand, the accessibility of the enzyme for organosoluble substrates is high when the interface is large, i.e., when the domain-size of the two phases is small. To investigate this, an amphiphilic network film which consisted of 75 wt % PHEA, 24 wt % poly(methyl acrylate), cross-linked by 1 wt % ethylene glycol dimethacrylate was prepared (referred to as PHEA-co-PMAA-lEGDMA). This network also shows a sponge-like morphology but with phase separation on a smaller scale than the PDMS cross-linked PHEA-l-PDMS-network (Figure 1d). The hydrophobic domains have diameters between 5 and 12 nm and the hydrophilic walls a thicknesses of 1 to 3.6 nm. The PHEA-l-PDMS-films were loaded with the enzyme horseradish peroxidase (HRP), which is known to catalyze synthetic relevant reactions such as polymerizations of electron-rich aromatics and asymmetric sulfoxidations. HRP’s catalytic activity in organic solvents is rather low,22-24 one reason being the enzyme’s insolubility in these media. The Nano Lett., Vol. 5, No. 1, 2005
Figure 2. (a) Microscope image of a film cross section of PHEA-l-PDMS and (b) fluorescence microscope image of this film loaded with fluorescence-labeled HRP.
films were immersed in an aqueous HRP solution [1 mg enzyme per mL buffer solution (100 mM phosphate, pH 7.0)] at 4 °C for 15 h, and afterward air-dried.25 The amount of enzyme within the coating was quantified spectrophotometrically by the Soret band of heme-containing proteins at 406 nm and was determined to be in the range of 10 to 24 µg cm-2 (5 to 12 µg HRP per mg polymer). The loading amount was confirmed by amino acid analysis. Upon storage at 4 °C, the biocatalysts did not lose any of its initial activity in water for at least 3 weeks. To address the question if the enzyme is within the film or just adsorbed to its surface, HRP was labeled with the fluorescence marker BODIPY FL (Molecular Probes) via its succinimidyl ester, following standard procedures.26 After loading the network with the labeled HRP, a cross section was investigated with a fluorescence microscope, revealing that the enzyme is indeed equally distributed within the whole network coating (Figure 2). The activity of the polymer-incorporated HRP in organic solvents was explored by means of the HRP-catalyzed oxidative coupling of N,N-dimethyl-p-phenylene diamine and phenol with tert-butyl hydroperoxide as model reaction (Figure 3), which gives in several steps an indophenol-type dye. Standard kinetic assays were not applicable.27 The specific catalytic activities were calculated from the linear slopes of the absorbance vs time plot. One Unit is defined as the increase in absorbance of 0.001 per minute at 546 nm at room temperature, with a reaction volume of 2.4 mL. The substrate concentrations of N,N-dimethyl-p-phenylene diamine and phenol were 1.67 mM, that of tert-butyl hydroperoxide was 0.83 mM. The linear slope indicates enzyme activity without deactivation. The length of it was taken for the operational stability of the peroxidases. The determined specific activities in n-heptane were in the range of 20 to 46 mU µg-1 when the peroxidase was encapsulated into the amphiphilic network and 0.44 ( 0.15 mU µg-1 for native HRP which was suspended directly in the reaction media, i.e., the activity was increased by 2 orders of magnitude. Not only the activity but also the operational stability were improved using the amphiphilic network as matrix for the enzyme (Figure 3). Altogether, the turnover of the matrixNano Lett., Vol. 5, No. 1, 2005
Figure 3. Peroxidase-catalyzed oxidative coupling reaction in n-heptane. Product formation of the reaction catalyzed by (9) 30 µg HRP in 1.3 cm2 of an PHEA-l-PDMS-film and (2) 750 µg of suspended native HRP, respectively. The conversion was monitored by the increase in absorbance (A) at 546 nm.
supported HRP was up to 230-fold higher after 80 min reaction time. Chemical reactions with immobilized enzymes may suffer from diffusion limitations.28,29 However, the monitored reaction shows a lag phase of about 25 min, independent of the kind of HRP used. This indicates that diffusional restrictions of substrates within the polymer play only a minor role for this conversion. Also, a lag phase is typical for this kind of reaction. The same reaction in a polar solvent, 2-propanol, showed no increased catalytic activity, possibly because the PHEA phase swells in this solvent rather than the PDMS phase. To prove the concept to be more general, a second enzyme, chloroperoxidase (CPO), was loaded into the amphiphilic PHEA-l-PDMS network from its aqueous solution [0.7 mg enzyme per mL buffer solution (96 mM phosphate, pH 7.0)]. In heptane, it showed an activity of 30 to 53 mU µg-1, compared to 3.5 ( 1.6 mU µg-1 for unsupported CPO. Operational stabilities for matrix-sup47
ported CPO of up to 225 min were observed, whereas the lyophilized CPO was completely inactivated within 15 min. No catalytic activity in n-heptane could be observed when HRP was loaded into nonamphiphilic polymer films consisting of 99 wt % PHEA and 1 wt % low molecular weight cross-linker ethyleneglycol dimethacrylate. The same observation was made for an HRP-loaded amphiphilic PHEA-coPMAA-l-EGDMA-film (morphology see Figure 1d). These findings show, that both the amphiphilic character and a nanophase-separated morphology with a minimal domain size in the dimension of the used enzyme (molecular radius HRP ) 3.0 nm 30) are crucial for the enhancement of its catalytic properties in nonpolar organic solvent. In conclusion, we have demonstrated for the first time that nanophase-separated amphiphilic networks possess the ability to stabilize and enhance the catalytic activity of enzymes in organic solvents. This is a new application for these materials and could broaden the applications of biocatalysts in organic synthesis on the lab scale as well as on an industrial scale. We expect that the catalytic properties of enzymes incorporated into an amphiphilic network can be increased further by optimizing the domain size of the network and by varying the chemical nature of the monomers. Future work will concentrate on other classes of enzymes and on synthetic relevant reactions such as chiral epoxidations, which are known to be catalyzed by peroxidases.22-24 Acknowledgment. This work was supported by the Deutsche Forschungsgemeinschaft (Emmy-Noether-Programm and SFB 428) and by the Fonds der Chemischen Industrie. We thank Dr. J. Scherble (Ro¨hm AG) for providing the macromonomers, Dr. R. Thomann for his help with the AFM measurements, and Prof. Dr. P. Gra¨ber (Freiburg) for help with the fluorescence labeling. Supporting Information Available: Experimental details of the synthesis of the amphiphilic networks, their loading with enzyme, and the kinetic assays. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Zaks, A.; Dodds, D. R. Drug DiscoV. Today 1997, 2(12), 513-531. (2) Rouhi, A. M. Chem. Eng. News 2003, 81(7), 55-73. (3) Klibanov, A. M. Nature 2001, 409, 241-246.
48
(4) Carrea, G.; Riva, S. Angew. Chem., Int. Ed. Engl. 2000, 39(13), 2226-2254. (5) Carrea, G.; Riva, S. Angew. Chem. 2000, 112(13), 2312-2341. (6) Schmid, A.; Dordick, J. S.; Hauer, B.; Kiener, A.; Wubbolts, M.; Witholt, B. Nature 2001, 409, 258-268. (7) Klibanov, A. M. Trends Biotechnol. 1997, 15, 97-101. (8) Burton, S. G.; Cowan, D. A.; Woodley, J. M. Nature Biotechnol. 2002, 20, 37-45. (9) Dai, L.; Klibanov, A. M. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 9475-9478. (10) Paradkar, V. M.; Dordick, J. S. J. Am. Chem. Soc. 1994, 116, 50095010. (11) Mabrouk, P. A. The use of poly(ethylene glycol) enzymes in nonaqueous enzymology. In Poly(ethylene glycol) Chemistry and Biological Applications; Harris, J. M., Zalipky, S., Eds.; American Chemical Society: Washington, DC, 1997; pp 118-133. (12) Wang, P.; Sergeeva, M. V.; Lim, L.; Dordick, J. S. Nature Biotechnol. 1997, 15, 789-793. (13) Yang, Z.; Mesiano, A. J.; Venkatasubramanian, S.; Gross, S. H.; Harris, J. M.; Russell, A. J. J. Am. Chem. Soc. 1995, 117, 48434850. (14) May, S. W. Curr. Opin. Biotechnol. 1997, 8, 181-186. (15) Ansorge-Schumacher, M. B.; Doumeche, B.; Metrangolo, D.; Hartmeier, W. MinerVa Biotechnol. 2000, 12(4), 265-359. (16) Gill, I.; Ballesteros, A. Trends Biotechnol. 2000, 18, 282-296. (17) Reetz, M. T. AdV. Mater. 1997, 9(12), 943-954. (18) Scherble, J., Ph.D. Thesis, Albert-Ludwigs-Universita¨t Freiburg (Germany), 2002. (19) Ivan, B.; Kennedy, J. P.; Mackey, P. W. Polym. Prepr. (Am. Chem. Soc., DiV. Polym. Chem.) 1990, 31(2), 217-218. (20) Partickios, C. S.; Georgiou, T. K. Curr. Opin. Colloid Interface Sci. 2003, 8, 76-85. (21) Soni, S.; Desai, J. D.; Devi, S. J. Appl. Polym. Sci. 2000, 77(13), 2996-3002. (22) Adam, W.; Lazarus, M.; Saha-Mo¨ller, C. R.; Weichold, O.; Hoch, U.; Ha¨ring, D.; Schreier, P. AdV. Biochem. Eng. Biotechnol. 1999, 63, 73-108. (23) van Deurzen, M. P. J.; van Rantwijk, F.; Sheldon, R. A. Tetrahedron 1997, 53(39), 13183-13220. (24) Colonna, S.; Gaggero, N.; Richelmi, C.; Pasta, P. Trends Biotechnol. 1999, 17, 163-168. (25) There was no detectable water-leaching from the dried films when immersed into heptane as determined by Karl Fischer titration. (26) Brinkley, M. Bioconjugate Chem. 1992, 3, 2-13. (27) Substrates (e.g., ABTS) and/or products (e.g., Oligoguaiacol) are not soluble in heptane. (28) Tischer, W.; Wedekind, F. Top. Curr. Chem. 1999, 200, 95-126. (29) Drauz, K.; Waldmann, H., Immobilization of Enzymes. In Enzyme Catalysis in Organic Chemistry, VCH: Weinheim, 1995; Vol. 1, pp 73-87. (30) Rennke, H. G.; Venkatachalam, M. A. J. Histochem. Cytochem. 1979, 27(10), 2352-1353.
NL048413B
Nano Lett., Vol. 5, No. 1, 2005
