Enzyme Stabilization and Immobilization: Methods and Protocols (Methods in Molecular Biology 679)

Methods

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Molecular Biology™

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Enzyme Stabilization and Immobilization Methods and Protocols

Edited by

Shelley D. Minteer Department of Chemistry, Saint Louis University, St. Louis, MO, USA

Editor Shelley D. Minteer, Ph.D. Department of Chemistry Saint Louis University St. Louis, MO USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-894-2 e-ISBN 978-1-60761-895-9 DOI 10.1007/978-1-60761-895-9 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010936521 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or ­dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, ­neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Enzyme stabilization has been an area of research interest since the 1950s, but in the last decade, researchers have made tremendous progress in the field. This has opened up new opportunities for enzymes in molecular biology as well as industrial applications, such as bioprocessing. The first chapter introduces the reader to the field of enzyme stabilization and the different theories of enzyme stabilization, including the use of immobilization as a stabilization technique. The first half of the book will focus on protocols for enzyme stabilization in solutions including liposome formation, micelle introduction, crosslinking, and additives, while the second half of the book will focus on protocols for enzyme stabilization during enzyme immobilization including common techniques like sol–gel encapsulation, polymer encapsulation, and single enzyme nanoparticle formation. Protocols for a variety of enzymes are shown, but the enzymes are chosen as examples to show that these protocols can be used for both enzymes of biological importance as well as enzymes of industrial importance. The final chapter will detail spectroscopic protocols, methods, and assays for studying the effectiveness of the enzyme stabilization and immobilization strategies. The chapters of this volume should provide molecular biologists, biochemists, and biomedical and biochemical engineers with the state-of-the art technical information required to effectively stabilize their enzyme of interest in a variety of environments (i.e., harsh temperature, pH, or solvent conditions). St. Louis, MO, USA

Shelley D. Minteer

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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  1 Introduction to the Field of Enzyme Immobilization and Stabilization . . . . . . . . . Michael J. Moehlenbrock and Shelley D. Minteer   2 Stabilization of Enzymes Through Encapsulation in Liposomes . . . . . . . . . . . . . . Makoto Yoshimoto   3 Micellar Enzymology for Thermal, pH, and Solvent Stability . . . . . . . . . . . . . . . . Shelley D. Minteer   4 Lipase Activation and Stabilization in Room-Temperature Ionic Liquids . . . . . . . Joel L. Kaar   5 Nanoporous Silica Glass for the Immobilization of Interactive Enzyme Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andreas Buthe, Songtao Wu, and Ping Wang   6 Enzyme Stabilization and Immobilization by Sol-Gel Entrapment . . . . . . . . . . . . Allan E. David, Arthur J. Yang, and Nam Sun Wang   7 Nanoporous Gold for Enzyme Immobilization . . . . . . . . . . . . . . . . . . . . . . . . . . Keith J. Stine, Kenise Jefferson, and Olga V. Shulga   8 Enzyme Stabilization via Bio-templated Silicification Reactions . . . . . . . . . . . . . . Glenn R. Johnson and Heather R. Luckarift   9 The Immobilization of Enzyme on Eupergit® Supports by Covalent Attachment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zorica D. Knežević-Jugović, Dejan I. Bezbradica, Dušan Ž. Mijin, and Mirjana G. Antov 10 Micellar Polymer Encapsulation of Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sabina Besic and Shelley D. Minteer 11 Enzyme Stabilization via Cross-Linked Enzyme Aggregates . . . . . . . . . . . . . . . . . Munishwar N. Gupta and Smita Raghava 12 Enzyme Immobilization in Polyelectrolyte Microcapsules . . . . . . . . . . . . . . . . . . . Michael J. McShane 13 Macroporous Poly(GMA-co -EGDMA) for Enzyme Stabilization . . . . . . . . . . . . . . Nenad B. Milosavić and Radivoje M. Prodanović 14 Enzyme–Nanoparticle Conjugates for Biomedical Applications . . . . . . . . . . . . . . Alexey A. Vertegel, Vladimir Reukov, and Victor Maximov 15 Enzyme Nanoparticle Fabrication: Magnetic Nanoparticle Synthesis and Enzyme Immobilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Patrick A. Johnson, Hee Joon Park, and Ashley J. Driscoll

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16 Cytochrome c Stabilization and Immobilization in Aerogels . . . . . . . . . . . . . . . . . 193 Amanda S. Harper-Leatherman, Jean Marie Wallace, and Debra R. Rolison 17 Kinetic Measurements for Enzyme Immobilization . . . . . . . . . . . . . . . . . . . . . . . 207 Michael J. Cooney Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227

Contributors Mirjana G. Antov  •  University of Novi Sad, Faculty of Technology, Novi Sad, Serbia Sabina Besic  •  Department of Chemistry, Saint Louis University, St. Louis, MO, USA Dejan I. Bezbradica  •  Department of Biochemical Engineering and Biotechnology, Faculty of Technology and Metallurgy, University of Belgrade, Belgrade, Serbia Andreas Buthe  •  Department of Bioproducts and Biosystems Engineering, University of Minnesota, St. Paul, MN, USA Michael J. Cooney  •  School of Ocean and Earth Science and Technology, Hawaii Natural Energy Institute, University of Hawaii-Manoa, Honolulu, HI, USA Allan E. David   •  Department of Chemical and Biomolecular Engineering, University of Maryland, College Park, MD, USA; Industrial Science & Technology Network (ISTN) Inc., York, PA, USA Ashley J. Driscoll  •  Department of Chemical and Petroleum Engineering, University of Wyoming, Laramie, WY, USA Munishwar N. Gupta  •  Chemistry Department, Indian Institute of Technology Delhi, Hauz Khas, New Delhi, India Amanda S. Harper-Leatherman  •  Chemistry Department, Fairfield University, Fairfield, CT, USA Kenise Jefferson  •  Department of Chemistry and Biochemistry, Center for Nanoscience, One University Boulevard, University of Missouri – Saint Louis, Saint Louis, MO, USA Glenn R. Johnson  •  Microbiology and Applied Biochemistry, Air Force Research Laboratory, Tyndall Air Force Base, FL, USA Patrick A. Johnson  •  Department of Chemical and Petroleum Engineering, University of Wyoming, Laramie, WY, USA Joel L. Kaar  •  Department of Chemical Engineering, McGowan Institute for Regenerative Medicine, University of Pittsburg, Pittsburg, PA, USA Zorica D. Kneževic´-Jugovic´  •  Department of Biochemical Engineering and Biotechnology, University of Belgrade, Belgrade, Serbia Heather R. Luckarift  •  Microbiology and Applied Biochemistry, Air Force Research Laboratory, Tyndall Air Force Base, FL, USA Victor Maximov  •  Department of Bioengineering, Clemson University, Clemson, SC, USA Michael J. McShane  •  Biomedical Engineering Department, Texas A&M University, College Station, TX, USA Dušan Ž. Mijin  •  University of Belgrade, Belgrade, Serbia Nenad B. Milosavic´  •  Department of Chemistry, Belgrade University, Belgrade, Serbia Shelley D. Minteer  •  Department of Chemistry, Saint Louis University, St. Louis, MO, USA

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Michael J. Moehlenbrock  •  Department of Chemistry, Saint Louis University, St. Louis, MO, USA Hee Joon Park  •  Department of Chemical and Petroleum Engineering, University of Wyoming, Laramie, WY, USA Radivoje M. Prodanovic´  •  University of Belgrade, Belgrade, Serbia Smita Raghava  •  Chemistry Department, Indian Institute of Technology Delhi, Hauz Khas, New Delhi, India Vladimir Rekov  •  Department of Bioengineering, Clemson University, Clemson, SC, USA Debra R. Rolison  •  Naval Research Laboratory, Surface Chemistry Branch, Washington, DC, USA Olga V. Shulga  •  Cross-Link Technologies, Jordan Valley Innovation Center, Springfield, MO, USA Keith J. Stine  •  Department of Chemistry and Biochemistry, University of Missouri – Saint Louis, Saint Louis, MO, USA Alexey A. Vertegel  •  Department of Bioengineering, Clemson University, Clemson, SC, USA Jean Marie Wallace  •  Nova Research, Inc., Alexandria, VA, USA Nam Sun Wang  •  Department of Chemical and Biochemical Engineering, University of Maryland, College Park, MD, USA Ping Wang  •  Department of Bioproducts and Biosystems Engineering, University of Minnesota, St. Paul, MN, USA Songtao Wu  •  Department of Bioproducts and Biosystems Engineering, University of Minnesota, St. Paul, MN, USA Arthur J. Yang  •  Industrial Science & Technology Network (ISTN) Inc., York, PA, USA Makato Yoshimoto  •  Department of Applied Molecular Bioscience, Yamaguchi University, Ube, Japan

Chapter 1 Introduction to the Field of Enzyme Immobilization and Stabilization Michael J. Moehlenbrock and Shelley D. Minteer Abstract Enzyme stabilization is important for any biomedical or industrial application of enzymes. In many applications, the goal is to provide extended active lifetime at normal environmental conditions with traditional substrates at low concentrations in buffered solutions. However, as enzymes are used for more and more applications, there is a desire to use them in extreme environmental conditions (i.e., high temperatures), in high substrate concentration, and in nontraditional solvent systems. This chapter introduces the topic of enzyme stabilization and the methods used for enzyme stabilization including enzyme immobilization. Key words: Enzyme immobilization, Enzyme stabilization, Crosslinking, Entrapment, Encapsulation

1. Introduction This book will detail methods for enzyme stabilization and enzyme immobilization. Although the concepts of enzyme immobilization and enzyme stabilization are different, they are related. Enzyme immobilization techniques are strategies focused on retaining an enzyme on a surface or support. In theory, they are focused on being able to reuse the enzyme and/or constrain the enzyme to a particular area of a reactor or device, whereas enzyme stabilization is focused on extending the active catalytic lifetime of the protein. Enzyme immobilization is one method for stabilizing enzymes. Enzymes are proteins with complex and fragile three-dimensional structures. The main methods for stabilizing enzymes are: protein engineering, chemical modification, immobilization, and adding additives for stabilization.

Shelley D. Minteer (ed.), Enzyme Stabilization and Immobilization: Methods and Protocols, Methods in Molecular Biology, vol. 679, DOI 10.1007/978-1-60761-895-9_1, © Springer Science+Business Media, LLC 2011

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Protein engineering is a very common method for enzyme stabilization, but it is out of the scope of this book, because it is not a simple or translational method that can be detailed in a chapter. The common types of protein engineering are directed evolution, site-directed mutagenesis, and peptide chain extensions. Directed evolution is a common method for enzyme engineering that involves a simple three step process that can be repeated as many “rounds”/repetitions as desired to evolve the mutant protein to have nonnatural attributes (in this case some type of stability). The three step process involves developing a library of random mutants, testing the library of mutants for the particular property/ attribute of interest, selecting the mutant of interest, and then repeating the above steps until optimized. Directed evolution will only be as good as the screening assay that you employ to test the library of mutants for the particular property/attribute of interest. This is frequently done for improving the temperature stability of proteins (1). Site-direct mutagenesis is another common technique that requires more understanding about the structure of the protein (2). Rather than random mutations, sitedirected mutagenesis involves an examination of the protein structure and intelligently mutating individual sites on the protein structure and then studying the effect of the site mutation on the particular property, activity, or attribute of interest (in this case stability). Peptide chain extensions are not a common technique, in general, for protein or enzyme engineering, but it is a technique for improving stability. Peptide chain extensions are the process of elongating the polypeptide chain of the enzyme on either the N terminus or the C terminus end. This has been shown to improve temperature stability of proteins (3, 4). It is not frequently done for biotechnology applications other than improving stability, but it can be effective.

2. Enzyme Stabilization This book will cover the other three methods of enzyme stabilization. The first half of the book will focus on chemical modification and adding additives for stabilization and the second half of the text will focus on enzyme immobilization. Chemical modification is common. The most common method is crosslinking. Crosslinkers are added for intramolecular crosslinking of the peptide chain. This is traditionally done with glutaraldehyde, but other bifunctional reagents can be used, including: dimethyl suberimidate, disuccinimidyl tartrate, and 1-ethyl-3-(3-dimethylaminopropryl) carbodiimide. It is important to note that this is unspecific chemistry as shown in Fig.  1 and therefore can cause both

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Introduction to the Field of Enzyme Immobilization and Stabilization

NaCNBH3

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Fig. 1. Options for glutaraldehyde crosslinking chemistry.

intramolecular and intermolecular crosslinking of the peptide chain, which can cause aggregation. This aggregation can be considered advantageous or disadvantageous depending on the application. Conjugation to water-soluble polymers like poly(ethylene glycol) (PEG) have been shown to be effective at stabilizing enzymes (5–8). This is frequently referred to as PEGylating the enzyme. PEGylating reagents contain a reactive function group either at one end of the PEG chain or at both ends of the PEG chain (bifunctional). If it is important to reduce intermolecular aggregation, monofunctional PEGylating reagents are typically used with functional groups to either react to the N terminus or C terminus end of the protein or to react with individual amino acids, such as: aspartic acid, glutamic acid, lysine, tyrosine, cysteine, histidine, and arginine. Other biocompatible polymers can also be employed, such as: chitosan (9), alginate, pectin, cellulose, hyaluronate, carrageenan, and poly(glutamate). These are becoming more popular in literature in recent years. The addition of additives to improve enzyme stabilization is also common. There are four main classes of additives: small molecules, polymer, proteins, and surfactant/micelles. The most common small molecule additive is trehalose. Trehalose is a disaccharide of two glucose units that is used for a variety of biological applications requiring dehydration and rehydration. For this reason, trehalose is a common additive when lyophilizing or freeze drying proteins. The trehalose sugar produces a gel as it dehydrates,

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thereby, protecting proteins, enzymes, organelles, cells, and tissue structure. Many sugars and sugar alcohols have also been employed for improving long-term storage and temperature stability, inclu­ ding: glycerol, lactose, mannitol, sorbitol, and sucrose. These are frequently referred to as lyophilization agents, because they improve storage stability during temperature fluctuations associated with freezing and thawing. Finally, polymers and proteins, such as: BSA, human albumin, gelatin, and PEG, have also been employed to improve long-term storage and/or lyophilization. Surfactants are another class of small molecules that are frequently added as additives to increase stability. Proteins have a tendency to denature at interfaces, so surfactants compete for binding at the interface. Also, it has been proposed that surfactants can facilitate refolding of partially denatured proteins. Micelles are also employed. Micelles are particularly useful for membrane-bound proteins, because the amphiphilic nature of the micelle and protein can be matched as well as providing an environment that is much more similar to cellular membranes than the buffer environment (10). This extends the active lifetime of the protein as well as being beneficial for stabilizing the protein in temperature environments, harsh pH environments, and organic solvents (11, 12).

3. Enzyme Immobilization Enzyme immobilization is a much broader topic that is arguably part science and part art (13). There are a number of factors to consider when choosing an enzyme immobilization strategy inclu­ ding: enzyme tolerance to immobilization, chemical and physical environment, surface functional groups on the protein, size of enzyme, charge on protein/pI, polarity of the protein (hydrophobic/ hydrophilic regions), and substrate/product transport needs (14). Each of these properties relates to different strategies for enzyme immobilization. The main types of enzyme immobilization are adsorption, crosslinking, and entrapment/encapsulation. Adsorption is the process of intermolecular forces resulting in the accumulation of protein on a solid surface. Adsorption is very dependent on the intermolecular interactions between the support surface and the enzyme. Therefore, properties such as enzyme charge and polarity are crucially important to ensure high and reproducible coverage of enzyme on the support. This is typically a mild immobilization technique (15). However, it is often problematic in that you typically have leaching of enzyme from the surface as a function of time. Covalent binding and crosslinking are techniques that allow for the covalent binding of enzyme to surfaces or to other enzymes. These surfaces could be inner walls of a bioreactor or

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supports like glass or polymeric beads for a packed-bed reactor for industrial bioprocessing. Covalent binding or crosslinking of enzyme to enzyme to form aggregates is commonly referred to as CLEAs (crosslinked enzyme aggregates) (16) and they have been used in a number of applications. The crosslinking of protein to protein or protein to surface typically results in decreasing the enzyme activity of the protein, but it provides a great deal of stability to the protein. Covalent binding and crosslinking chemistry can sometimes be harsh chemistry. Therefore, it is important to consider the chemical microenvironment of the chemistry before use with proteins or enzymes, due to the fragility of the protein three-dimensional structure. The final method of enzyme immobilization is entrapment or encapsulation within a polymeric matrix. Entrapment is the procedure of polymerizing a monomer or low molecular weight polymer around the protein to trap the protein on a surface. This is frequently done with sol-gels and hydrogels and is quite successful at immobilizing proteins on surfaces (17–20). Traditional and electropolymerized polymers can also be employed for entrapping enzymes (21, 22). Issues to consider with entrapment techniques are the chemical environment of the polymerization solution and whether it will denature the protein as well as the pore size and interconnectivity of the pores in the polymer to determine if transport of substrate and product can diffuse in and out of the polymer, but to ensure that the protein cannot diffuse out of the polymers. Polymer entrapment has been common, because many strategies do not result in a significant decrease in enzyme activity after immobilization. However, leaching is frequently a problem depending on enzyme loading and the entrapment technique tends to have less of an effect on enzyme stabilization in high temperatures and organic solvents than crosslinking techniques. Encapsulation is similar to entrapment in that the protein is constrained within the polymeric matrix, but the polymer matrix has “pockets” or “pores” for immobilizing the protein. Micellar polymers are an example of polymers that can encapsulate an enzyme. The polymer micelles are swelled and the enzyme is allowed to intercalate into the micellar pockets/aggregates and the solvent is evaporated. This provides a polymer membrane that provides a microenvironment similar to a cellular microenvironment that can provide temperature, pH, and organic solvent stability to the protein (23).

4. Applications Immobilized and stabilized enzymes have a variety of applications. Stabilization is important for the commercialization of industrial enzymes, because the enzymes must be stabilized for

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shipping and storage. Therefore, the small molecule additives are common for the commercial production and sale of enzymes of all quantities. However, stabilization is more difficult in industrial settings, so bioprocessing applications requiring enzymes to be immobilized on the inside of reactors or on supports in a packed bed reactor are typically more challenging than just stabilizing an enzyme for distribution and storage. Bioprocessing applications range from food applications (i.e., the use of glucose isomerase in the production of corn syrup) to pharmaceutical applications (i.e., enzyme to ensure chirality of drug products) to fuel applications (i.e., enzymatic production of butanol). Other common applications of immobilized enzymes are biosensors and biofuel cells. Biosensors employ enzymes to selectively catalyze the reaction of an analyte (24). Biofuel cells employ enzymes to catalyze the reduction of oxygen at the cathode of a fuel cell and/or the oxidation of fuel at the anode of a fuel cell (25). The hostile pH and electric field environment in these applications result in a need for enzyme stabilization as well as immobilization on the electrode surface to improve efficiency. Overall, there are a wealth of industrial and laboratory applications for immobilized and stabilized enzymes. The following chapters will give protocols and examples for different strategies and techniques for immobilization and stabilization. References 1. Hecky, J., and Muller, K. M. (2005) Structural perturbation and compensation by directed evolution at physiological temperature leads to thermostabilization of beta-lactamase, Biochemistry 44, 12640–12654. 2. O’Fagain, C. (2003) Enzyme stabilization – recent experimental progress, Enzyme and Microbial Technology 33, 137–149. 3. Liuu, J. H., Tsai, F. F., Liu, J. W., Cheng, K. J., and Cheng, C. L. (2001) The catalytic domain of a Piromyces rhizinflata cellulase expressed in E. coli was stabilized by the linker peptide of the enzyme, Enzyme and Microbial Technology 28, 582–589. 4. Matsura, T., Miyai, K., Trakulnaleamsai, S., Yomo, T., Shima, Y., Miki, S., Yamamoto, K., and Urabe, I. (1999) Evolutionary molecular engineering by random elongation mutagenesis, Nature Biotechnology 17, 58–61. 5. Jeng, F. Y., and Lin, S. C. (2006) Characterization and application of PEGylated horseradish peroxidase for the synthesis of poly(2-naphthol), Process Biochemistry 41, 1566–1573. 6. Treetharnmathurot, B., Ovartlarnporn, C., Wungsintaweekul, J., Duncan, R., and

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Wiwattanapatapee, R. (2008) Effect of PEG molecular weight and linking chemistry on the biological activity and thermal stability of PEGylated trypsin, International Journal of Pharmaceutics 357, 252–259. Veronese, F. M. (2001) Peptide and protein PEGylation: a review of problems and solutions, Biomaterials 22, 405–417. Roberts, M. J., Bentley, M. D., and Harris, J. M. (2002) Chemistry of peptide and protein PEGylation, Advanced Drug Delivery Reviews 54, 459–476. Gomez, L., Ramırez, H. L., Villalonga, M. L., Hernandez, J., and Villalonga, R. (2006) Immobilization of chitosan-modified invertase on alginate-coated chitin support via polyelectrolyte complex formation, Enzyme and Microbial Technology 38, 22–27. Martinek, K., Klyachko, N. L., Kabanov, A. V., Khmel’nitskii, Y. L., and Levashov, A. V. (1989) Micellar enzymology: its relation to membranology, Biochimica et Biophysica Acta 981, 161–172. Martinek, K., Klyachko, N. L., Levashov, A. V., and Berezin , I. V. (1983) Micellar enzymology.

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13. 14. 15.

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17.

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Catalytic activity of peroxidase in a colloidal aqueous solution in an organic solvent, Doklady Akademii Nauk SSSR 263, 491–493. Celej, M. S., D’Andrea, M. G., Campana, P. T., Fidelio, G. D., and Bianconi, M. L. (2004) Superactivity and conformational changes on chymotrypsin upon interfacial binding to cationic micelles, Biochemical Journal 378, 1059–1066. Cao, L. (2005) Immobilised enzymes: science or art?, Current Opinion in Chemical Biology 9, 217–226. Hanefeld, U., Gardossi, L., and Magner, E. (2009) Understanding enzyme immobilisation, Chemical Society Reviews 38, 453–468. Cooney, M. J., Svoboda, V., Lau, C., Martin, G. P., and Minteer, S. D. (2008) Enzyme catalysed biofuel cells, Energy & Environmental Science 1, 320–337. Kim, M. I., Kim, J., Lee, J., Jia, H., Na, H. B., Youn, J. K., Kwak, J. H., Dohnalkova, A., Grate, J. W., Wang, P., Hyeon, T., Park, H. G., and Chang, H. M. (2006) Crosslinked enzyme aggregates in hierarchically ordered mesoporous silica: a simple and effective method for enzyme stabilization, Biotechnology and Bioengineering 96, 210–218. Coche-Guerente, L., Cosnier, S., and Labbe, P. (1997) Sol-gel derived composite materials for the construction of oxidase/peroxidase mediatorless biosensors, Chemistry of Materials 9, 1348–1352. Lim, J., Malati, P., Bonet, F., and Dunn, B. (2007) Nanostructured sol-gel electrodes for

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22.

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24. 25.

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biofuel cells, Journal of the Electrochemical Society 154, A140–A145. Nguyen, D. T., Smit, M., Dunn, B., and Zink, J. I. (2002) Stabilization of creatine kinase encapsulated in silicate sol-gel materials and unusual temperature effects on its activity, Chemistry of Materials 14, 4300–4306. Hussain, F., Birch, D. J. S., and Pickup, J. C. (2005) Glucose sensing based on the intrinsic fluorescence of sol-gel immobilized yeast hexokinase, Analytical Biochemistry 339, 137–143. Yang, R., Ruan, Y., and Deng, J. (1998) A H2O2 biosensor based on immobilization of horseradish peroxidase in electropolymerized methylene green film on GCE, Journal of Applied Electrochemistry 28, 1269–1275. Chiang, C.-J., Hsiau, L.-T., and Lee, W.-C. (2004) Immobilization of cell-associated enzymes by entrapment in polymethacrylamide beads, Biotechnology Techniques 11, 121–125. Moore, C. M., Akers, N. L., Hill, A. D., Johnson, Z. C., and Minteer, S. D. (2004) Improving the environment for immobilized dehydrogenase enzymes by modifying Nafion with tetraalkylammonium bromides, Biomacro­ molecules 5, 1241–1247. Aston, W. J., and Turner, A. P. F. (1984) Biosensors and biofuel cells, Biotechnology and Genetic Engineering Reviews 1, 89–120. Atanassov, P., Apblett, C., Banta, S., Brozik, S., Calabrese-Barton, S., Cooney, M. J., Liaw, B. Y., Mukerjee, S., and Minteer, S. D. (2007) Enzymatic biofuel cells, Electrochemical Society Interface 16, 28–31.

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Chapter 2 Stabilization of Enzymes Through Encapsulation in Liposomes Makoto Yoshimoto Abstract Phospholipid vesicle (liposome) offers an aqueous compartment surrounded by lipid bilayer membranes. Various enzyme molecules were reported to be encapsulated in liposomes. The liposomal enzyme shows peculiar catalytic activity and selectivity to the substrate in the bulk liquid, which are predominantly derived from the substrate permeation resistance through the membrane. We reported that the quaternary structure of bovine liver catalase and alcohol dehydrogenase was stabilized in liposomes through their interaction with lipid membranes. The method and condition for preparing the enzyme-containing liposomes with well-defined size, lipid composition, and enzyme content are of particular importance, because these properties dominate the catalytic performance and stability of the liposomal enzymes. Key words: Liposomes, Phospholipid vesicles, Lipid bilayer membranes, Enzyme encapsulation, Membrane permeability, Enzyme structure, Enzyme reactivity, Bovine liver catalase

1. Introduction The liposomal aqueous phase is isolated from the bulk liquid by the semipermeable lipid bilayer membranes, which means chemical reactions can be induced inside enzyme-containing liposomes by adding membrane-permeable substrate to the bulk liquid. In the liposomal system, the enzyme molecules are confined without chemical modification, which is advantageous to preserve the inherent enzyme affinity to the cofactor and substrate molecules. So far, various liposome-encapsulated enzymes have been prepared and characterized for developing diagnostic and biosensing materials, functional drugs, and biocompatible catalysts (1, 2). The reactivity of liposomal enzymes was extensively examined mainly focusing on the membrane permeation of the substrate molecules as a rate-controlling step of the liposomal reaction (3, 4). For example, sodium Shelley D. Minteer (ed.), Enzyme Stabilization and Immobilization: Methods and Protocols, Methods in Molecular Biology, vol. 679, DOI 10.1007/978-1-60761-895-9_2, © Springer Science+Business Media, LLC 2011

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cholate is a useful modulator of the liposome membranes. Incorporation of sublytic concentrations of cholate in the membranes induced permeation of substrate and as a result the rate of liposomal enzyme reaction increased (4). An excess amount of cholate causes complete solubilization of liposome membrane, which is utilized for determining the total amount and inherent activity of the enzyme encapsulated in liposomes. On the other hand, the stability of enzyme activity in liposomes is relatively unknown. We recently reported that the thermostability of bovine liver catalase and yeast alcohol dehydrogenase consi­derably increased through encapsulation of each enzyme in liposomes composed of POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine) (5–7). Furthermore, the liposomal glucose oxidase system was shown to be applicable as a stable catalyst for the prolonged oxidation of glucose in the gas–liquid flow in a bubble column reactor (8–10). In the liposomal system, the aggregate formation among the partially denatured enzyme molecules was indicated to be depressed through the interaction of the enzyme with lipid membranes (11). This chapter describes the preparation, reactivity, and stability of the enzyme-containing liposomes with various sizes and enzyme contents using the catalase as a model enzyme. The preparation and analytical methods described are basically applicable to liposomes containing other water-soluble enzymes. To prepare stable and reactive liposomal enzyme systems, the enzyme content in liposomes and the lipid composition need to be changed and optimized considering the characteristics of each enzyme employed and the permeability of its substrate through the liposome membranes.

2. Materials 2.1. Preparation of Catalase-Containing Liposomes

1. Phospholipid: POPC (>99%, Mr = 760.1, main phase transition temperature Tm of −2.5 ± 2.4°C (12)) (Avanti Polar Lipids, Inc., Alabaster, AL). 2. Chloroform (>99%). Diethylether (>99.5%). 3. Ethanol (>99.5%). 4. Dry ice (solid CO2). 5. Rotary evaporator (REN-1, Iwaki Co., Ltd., Japan) with an aspirator (ASP-13, Iwaki Co., Ltd.). 6. Freeze-dryer (FRD-50  M, Asahi Techno Glass Corp., Funabashi, Japan) with a vacuum pump (GLD-051, ULVAC, Inc., Chigasaki, Japan). 7. Enzyme: bovine liver catalase (EC 1.11.1.6, ca. 10,000 U/mg, Mr = 240,000) (Wako Pure Chemical Industries, Ltd., Osaka, Japan).

Stabilization of Enzymes Through Encapsulation in Liposomes

11

8. Tris buffer: 50  mM Tris (2-amino-2-hydroxymethyl-1, 3-propanediol)–HCl, pH 7.4, containing 0.1  M sodium chloride. 9. Small-volume extrusion device Liposofast™ and its stabilizer (Avestin Inc., Ottawa, Canada) (13). 10. Polycarbonate membranes for sizing liposomes (Avestin, Inc., 19 mm in membrane diameter and 30, 50, or 100 nm in the nominal mean pore diameter). 11. Gel beads for gel permeation chromatography (GPC): sepharose 4B suspended in ethanol/water (GE Healthcare UK Ltd., Buckinghamshire, England). 12. Glass column with a stopcock for the GPC, 1.0 (id) × 35 cm, 20 mL in packed gel bed volume. 13. Enzyme kit for the quantification of POPC (Phospholipid C-Test Wako, Wako Pure Chemical Industries, Ltd.). 14. UV/visible spectrophotometer (Ubest V-550DS, JASCO, Tokyo, Japan) equipped with a perche-type temperature controller (EHC-477S, JASCO). 2.2. Measurement of Enzyme Activity of Catalase-Containing Liposomes

1. 0.3 M sodium cholate (Wako Pure Chemical Industries, Ltd.) in the Tris buffer. 2. Substrate of catalase: hydrogen peroxide (H2O2) solution (Wako Pure Chemical Industries, Ltd.).

3. Methods 3.1. Preparation of Catalase-Containing Liposomes

1. Weigh 50 mg of POPC powder (see Note 1). 2. Dissolve 50 mg of POPC in 4 mL of chloroform in a 100-mL round-bottom flask in a draft chamber. 3. Remove the solvent from the flask by using the rotary evaporator under reduced pressure in a draft chamber to form a lipid film on the inner wall of the flask. 4. Dissolve the lipid film in 4 mL of diethylether and remove the solvent as described above. Repeat this procedure once more. 5. Dry the lipid film formed in the flask by using the freezedryer connected to the vacuum pump for 2 h in the dark to remove the residual organic solvents molecules in the lipid layers. Keep the inner pressure of the flask 20 h). 8. To ensure the settings of the GC have not changed with time, which may alter the retention times and peak size of the reaction products, the GC should be calibrated daily by a single-point calibration; otherwise, an internal standard should be used. 9. It is best to do the titration against a white backdrop so as to make to the color of the titration solution easily identifiable. As the titration endpoint is neared, the solution may appear blue upon addition of the indicator dye only to become translucent after swirling. The titration should not be stopped until a blue tint remains after swirling. 10. Our results found that immobilized forms of lipase retained nearly full activity upon incubation in [bmim][PF6] over 24 h at 30°C (5). Immobilized lipase was considerably less stable in RTILs containing [bmim] and 1-methyl-1-(-2-methoxyethyl) pyrrolidinium ([mmep]) cations with the nitrate anion at the same temperature. Interestingly, incubation in select RTILs, including [mmep][CH3SO3], [bmim][CH3CO2], and [mmep][CH3CO2], resulted in a significant increase in lipase (Novozym 435) activity when returned to water. The increased activity of the immobilized lipase may be explained

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Kaar

by swelling of the immobilization support in these RTILs, causing previously inaccessible enzyme to be solvent exposed and a resultant increase in active site concentration. Though seemingly less likely, it is also plausible that the RTIL-exposed enzyme has a modified tertiary structure with increased activity in water.

Acknowledgments This work was funded by SACHEM Inc. and by a research grant from the Environmental Protection Agency (R-82813101-0) to Alan J. Russell, to whom I am grateful for support, both financial and scientific. I am thankful to Jason A. Berberich for technical advice and helpful discussion on all aspects of this work. I also wish to thank Jason A. Berberich and Rick R. Koepsel for their critical reading of this manuscript. References 1. Brennecke, J. F. and Maginn, E. J. (2001) Ionic liquids: innovative fluids for chemical processing, AIChE J 47, 2384–2389. 2. Yang, Z. and Pan, W. (2005) Ionic liquids: green solvents for nonaqueous biocatalysis, Enzyme Microb Technol 37, 19–28. 3. van Rantwijk, F. and Sheldon, R. A. (2007) Biocatalysis in ionic liquids, Chem Rev 107, 2757–2785. 4. Roosen, C., Muller, P., and Greiner, L. (2008) Ionic liquids in biotechnology: applications and perspectives for biotransformations, Appl Microbiol Biotechnol 81, 607–614. 5. Kaar, J. L., Jesionowski, A. M., Berberich, J. A., Moulton, R., and Russell, A. J. (2003) Impact of ionic liquid physical properties on lipase activity and stability, J Am Chem Soc 125, 4125–4131. 6. Micaêlo, N. M. and Soares, C. M. (2008) Protein structure and dynamics in ionic liquids. Insights from molecular dynamics simulation studies, J Phys Chem B 112, 2566–2572. 7. Halling, P. J. (1994) Thermodynamic predictions for biocatalysis in nonconventional media: theory, tests, and recommendations for experimental design and analysis, Enzyme Microb Technol 16, 178–206. 8. Halling, P. J. (2004) What can we learn by studying enzymes in non-aqueous media? Philos Trans R Soc Lond B Biol Sci 359, 1287–1296; discussion 1296–1287, 1323–1288.

9. Nakashima, K., Maruyama, T., Kamiya, N., and Goto, M. (2006) Homogeneous enzymatic reactions in ionic liquids with poly(ethylene glycol)-modified subtilisin, Org Biomol Chem 4, 3462–3467. 10. Berberich, J. A., Kaar, J. L., and Russell, A. J. (2003) Use of salt hydrate pairs to control water activity for enzyme catalysis in ionic liquids, Biotechnol Prog 19, 1029–1032. 11. Lejeune, K. E., Mesiano, A. J., Bower, S. B., Grimsley, J. K., Wild, J. R., and Russell, A. J. (1997) Dramatically stabilized phosphotriesterase-polymers for nerve agent degradation, Biotechnol Bioeng 54, 105–114. 12. Gordon, R. K., Feaster, S. R., Russell, A. J., LeJeune, K. E., Maxwell, D. M., Lenz, D. E., Ross, M., and Doctor, B. P. (1999) Organophosphate skin decontamination using immobilized enzymes, Chem Biol Interact 119-120, 463–470. 13. LeJeune, K. E., Swers, J. S., Hetro, A. D., Donahey, G. P., and Russell, A. J. (1999) Increasing the tolerance of organophosphorus hydrolase to bleach, Biotechnol Bioeng 64, 250–254. 14. Gill, I. and Ballesteros, A. (2000) Bioencap­ sulation within synthetic polymers (Part 2): non-sol-gel protein-polymer biocomposites, Trends Biotechnol 18, 469–479. 15. Drevon, G. F., Hartleib, J., Scharff, E., Ruterjans, H., and Russell, A. J. (2001)

Lipase Activation and Stabilization in Room-Temperature Ionic Liquids

16.

17.

18.

19.

20.

Thermoinactivation of diisopropylfluorophosphatase-containing polyurethane polymers, Biomacromolecules 2, 664–671. Drevon, G. F., Danielmeier, K., Federspiel, W., Stolz, D. B., Wicks, D. A., Yu, P. C., and Russell, A. J. (2002) High-activity enzymepolyurethane coatings, Biotechnol Bioeng 79, 785–794. Vasudevan, P. T., Lopez-Cortes, N., Caswell, H., Reyes-Duarte, D., Plou, F. J., Ballesteros, A., Como, K., and Thomson, T. (2004) A novel hydrophilic support, CoFoam, for enzyme immobilization, Biotechnol Lett 26, 473–477. Burrell, A. K., Del Sesto, R. E., Baker, S. N., McCleskey, T. M., and Baker, G. A. (2007) The large scale synthesis of pure imidazolium and pyrrolidinium ionic liquids, Green Chem 9, 449–454. Park, S. and Kazlauskas, R. J. (2001) Improved preparation and use of room-temperature ionic liquids in lipase-catalyzed enantio- and regioselective acylations, J Org Chem 66, 8395–8401. Lee, S. H., Ha, S. H., Lee, S. B., and Koo, Y. M. (2006) Adverse effect of chloride impurities on lipase-catalyzed transesterifications in ionic liquids, Biotechnol Lett 28, 1335–1339.

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21. Lejeune, K. E. and Russell, A. J. (1996) Covalent binding of a nerve agent hydrolyzing enzyme within polyurethane foams, Biotechnol Bioeng 51, 450–457. 22. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal Biochem 72, 248–254. 23. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Measurement of protein using bicinchoninic acid, Anal Biochem 150, 76–85. 24. Pace, C. N., Vajdos, F., Fee, L., Grimsley, G., and Gray, T. (1995) How to measure and predict the molar absorption coefficient of a protein, Protein Sci 4, 2411–2423. 25. Halling, P. J. (1992) Salt hydrates for water activity control with biocatalysts in organic media, Biotechnol Tech 6, 271–276. 26. Zacharis, E., Omar, I. C., Partridge, J., Robb, D. A., and Halling, P. J. (1997) Selection of salt hydrate pairs for use in water control in enzyme catalysis in organic solvents, Biotechnol Bioeng 55, 367–374.

w

Chapter 5 Nanoporous Silica Glass for the Immobilization of Interactive Enzyme Systems Andreas Buthe, Songtao Wu, and Ping Wang Abstract Recent pursuit on utilization of nanoscale materials has manifested a variety of configurations of highly efficient enzymic biocatalyst systems for biotechnological applications. Nanoscale structures are particularly powerful in effecting multienzyme biocatalysis. Inherent properties of nanomaterials – primarily, the high surface area to volume ratio and atomic scale 3D configurations – enable higher enzyme loadings, microenvironment control surrounding enzyme molecules, regulation on mass transfer, and protein structural stabilization of the biocatalyst as compared to traditional immobilization systems. This chapter introduces one versatile nanoscale immobilization method via details demonstrated using the case of nanoporous silica glass (30 nm diameter) for the concomitant incorporation of lactate dehydrogenase (LDH), glucose dehydrogenase (GDH), and the cofactor (NADH). Key words: Nanoporous carrier, Mesoporous silica glass, Covalent binding, Coupling agent, Cofactor regeneration, NADH, Lactate dehydrogenase, Glucose dehydrogenase

1. Introduction A number of methods can be used for the immobilization of an enzyme catalyst, but the choice of the right one is certainly a challenging task. Each method offers some advantages with regard to its effects on biocatalysts, but it has its drawbacks as well (1, 2). The literature provides little guidance on selection of proper methods for a certain application. Each enzyme and reaction has highly specific demands that cannot be fulfilled by following a simple standard procedure, especially if large-scale applications are involved. Typically, different methods need to be screened for their principle suitability, followed by the selection of the most promising method that has to be fine tuned to obtain the desired performance, admittedly a time-consuming procedure. As identified Shelley D. Minteer (ed.), Enzyme Stabilization and Immobilization: Methods and Protocols, Methods in Molecular Biology, vol. 679, DOI 10.1007/978-1-60761-895-9_5, © Springer Science+Business Media, LLC 2011

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by previous researchers, there are several key factors that influence the efficiency of an immobilized biocatalyst. These can be basically distinguished as those that affect the enzyme’s microenvironment and those that affect the mass transfer of substrate and product to and from the active site of the enzyme (2, 3). The presence of a favorable and protective microenvironment for the biocatalyst is a prerequisite for obtaining a high activity and stability; on the other hand, it may also invite a severe mass transfer limitation (1). Balancing between these tradeoffs is a key issue and becomes possible, e.g., by entrapping the enzymes in the nanopores of mesoporous silica glass by either physical adsorption or chemical binding. The highly specific surface area of these materials allows a high enzyme loading with an overwhelming majority of enzyme molecules accessible by the substrate; at the same time, the nanoscale structure of the porous support material protects the enzyme from detrimental effects and additionally exerts a beneficial impact on the enzyme stability (4–6). Thereby, the mechanism of enzyme stabilization is different from that involved in macroscopic materials normally used in enzyme immobilization. The entrapment in nanopores means confining the enzyme molecules in a space of comparable size, thus limiting the spatial volume available for the enzyme to unfold (5). A further improvement of the operational stability can be achieved by covalent bonding of the enzymes to the nanoporous material, which equally suppresses conformational changes and the leaching of the enzyme. Certainly, as the biocatalyst is confined to a limited space, this is likely associated with some mass transfer limitations, but to a lower extent than for other immobilization methods (e.g., if enzymes are entrapped in a polymer). Enabling the design of molecular machinery is another remarkable advantage of the entrapment in nanopores, as was demonstrated previously (7): if more than one enzyme is contained in such a pore, it becomes possible to catalyze multiple reactions, thus mimicking specific biological transformations. The objective of this chapter is to guide the reader through a protocol that aims at the coimmobilization of two enzymes, namely, lactate dehydrogenase (LDH) and glucose dehydrogenase (GDH) by incorporation into glass nanoporous materials with pores of 30 nm in diameter (Fig. 1). In order to avoid the loss of cofactor, the necessary NAD(H)molecules are immobilized through a flexible tether, by which they can shuttle back and forth between the two enzymes. The involved procedure comprises the activation of the porous glass support by an alkoxysilane and the subsequent covalent binding of the enzyme molecules and cofactor via a bifunctional agent. The procedure is easy to follow, requires no special equipment, and certainly offers a broad applicability, thus allowing the immobilization of any enzyme (see Note 1).

Nanoporous Silica Glass for the Immobilization of Interactive Enzyme Systems

a

b

Pyruvate

NADH

D-Glucono-1,5-lactone HO OH

OH O

O

HO

OH

HO HOH2C

LDH

O

GDH

HO

O

+

L-Lactate

HOH2C

O

OH

HO NAD

39

OH O

D-Glucose

Fig. 1. (a) Configuration of the immobilized multiple enzyme system with coimmobilized cofactor in the mesoporous silica glass with a pore diameter of 30 nm, enzymes and cofactor are covalently bound within the pores, (b) reaction scheme of the enzyme-coupled cofactor regeneration under the reduction of pyruvate to lactate catalyzed by lactate dehydrogenase (LDH) and the concomitant oxidation of glucose to gluconolactone catalyzed via glucose dehydrogenase.

2. Materials 2.1. Chemicals

●●

Porous silica glasses of surface area of 50–100 m2/g (Silicycle, Quebec, QC, Canada).

●●

Epichlorhydrin (ECH; Milwaukee, WI).

●●

3-Aminopropyltrimethoxysilane (APTMS; Milwaukee, WI).

●●

Ethanol (HPLC grade), sodium phosphate, glutardialdehyde (GDA), poly(ethyleneglycol) (PEG, MW 10,000 Da), Bradford reagent, bovine serum albumin (BSA), LDH, GDH, NADH, sodium pyruvate, glucose, glucose oxidase, horseradish peroxidase, o-dianisidine dihydrochloride are purchasable from Sigma-Aldrich Co. (St. Louis, MO).

2.2. Equipment

All equipment needed to apply this method should be available in a common laboratory, including: a UV-VIS spectrophotometer, a HPLC system, a pH probe, a horizontal shaker, a water bath, and a magnetic stirrer. As for the disposable labware, we recommend the use of microcentrifuge tubes (1.5–2.0 mL), 15-mL centrifuge tubes (see Note 2), screw-capped glass vials of 10–20 mL, and 0.2 mm PTFE syringe filters.

2.3. Solutions

The following provides sufficient amounts for the preparation of three batches of a 1-g immobilizate (see Note 3).

2.3.1. For Activation of Glass

1. Solution A: 100 mL 80:20 (v/v) ethanol-water. 2. Solution B: 25 mL of solution A + 2.5 g APTMS (0.05 g/mL). 3. Solution C: 25-mL phosphate buffer (50 mM/pH 7.0) + 1 g GDA (0.02 g/mL). 4. Solution D: 500-mL phosphate buffer (50 mM/pH 7.0). 5. Solution E (optional): 25-mL phosphate buffer (50  mM/ pH 7.0) + 30 mg/mL BSA.

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2.3.2. For Enzyme and Cofactor Attachment

1. Solution F: 25-mL solution D + 7.5 mg/mL NADH.

2.3.3. For Determination of Enzyme Activity

1. Solution H: 100 mL of solution D containing 1 mM pyruvate and 1 mM glucose.

2. Solution G: 25-mL solution D + 87.5  mg/mL LDH and 67.5 mg/mL GDH (see Note 4).

2. Solution I: (PGO-colorant solution): 100 mL of solution D with 5 U/mL glucose oxidase, 1 U/mL horseradish peroxidase, and 40 mg/mL o-dianisidine dihydrochloride. 3. Eluent for HPLC: 50:50 (v/v) acetonitrile–water, pH 2.1 (adjusted by sulfuric acid).

3. Methods 3.1. Activation of Nanoporous Glass

To ensure the highest possible benefit from the nanoscale immobilization in terms of biocatalytic activity, inert porous materials with a specific surface area from 50 to 100 m2/g and pore diameters up to 100 nm should be utilized. Those requirements can be best fulfilled by mesoporous silica glass, that can be purchased as bulk powder, as sintered glass (e.g., filled in a filter catridge), or even self-produced by common protocols (8, 9). The preferred source and shape depends on the targeted application. For simplicity, we recommend the use of commercially available mesoporous glass. To achieve a covalent bonding between the enzyme molecules and the glass support, the latter needs to be activated by silanization. Thereby, the inorganic glass carrier is exposed to an organosilane as a coupling agent, which contains at least one terminal alkoxy-silyl group and a terminal organic functionality (predominantly an amino group); however, epoxy, vinyl, or sulfhydryl groups can also be introduced for activation (10). Coupling of the silane to the carrier is achieved by the hydrolysis of the alkoxy group and the subsequent condensation between the silanol groups of the support material and the coupling agent (Fig. 2). Several protocols can be obtained from the literature, in which, for example, the silianization is conducted in either polar organic solvents or aqueous solution. Best results in our lab were obtained by utilizing APTMS for activation solubilized in an ethanol–water solution. OH

OC2H5

–O–Si–(CH2)3–NH2

–OH + H5C2O–Si–(CH2)3–NH2 OC2H5

C2H5OH

OH

Fig. 2. Chemical route for the activation of glass supports with 3-aminopropyltrimethoxysilane (APTMS).

Nanoporous Silica Glass for the Immobilization of Interactive Enzyme Systems

41

1. One gram (1 g) of the glass material is suspended in 8 mL of solution B and gently stirred for a duration of 2 h at 40°C (see Note 5). For an efficient activation of the mesoporous glass, the chosen duration and temperature are of importance to allow the complete formation of Si–O–Si bonds. 2. Activated glass is removed from solution by centrifugation (see Note 2), followed by washing with solution A at least three times to make sure that all unreacted organosilane is removed. 3. The activated mesoporous glass can be dried by air; even though further use in wet state is possible, it would hamper the mass balance due to the residual water. Storage under refrigeration is recommended. 3.2. Modification with Spacers

Sometimes, better results are obtained if a spacer between the support and the enzyme is used (7, 11–14). In particular, for redox-enzymes with a NAD(P)H cofactor requirement, it has been proven that the co-immobilization of the cofactor via a spacer heightens the overall catalytic efficiency, since the spacers allow the facilitated docking of the cofactor to the active site and its “traveling” between two redox-enzymes that are used for the cofactor regeneration (4). Therefore, we describe the further modification of the activated glass with spacers of different size (length) before the enzymes and cofactor are attached. Thereby, the spacers act simply as bifunctional coupling agents that link enzyme and activated support. The simplest spacer that can be used is glutardialdehyde (GDA), which reacts with amino groups, previously introduced by activation, under the formation of a Schiff base (Fig. 3).

3.2.1. Modification with GDA

1. One gram (1 g) of the previously activated glass is suspended in 8  mL of solution C and gently shaken for 4  h at room temperature (see Notes 5 and 6).

activated glass

2. Spacer-modified glass is removed from solution by centrifugation, followed by washing with solution D at least three times to make sure that all unreacted GDA is removed. OH –O –Si–(CH2)3–NH2 + OHC-CH2-CH2-CH2-CHO + H2N– E glutardialdehyde

OH OH

–O–Si–(CH2)3–N =CH-CH2-CH2-CH2-CH=N– E OH

Fig. 3. Coupling of enzyme and activated glass support through the bifunctional agent glutardialdehyde (GDA) under the formation of Schiff base.

42

Buthe, Wu, and Wang O H2C–CH–CH2–Cl +

2

epichlorohydrin

HO–CH2–CH2–O–CH2–CH2–OH poly(ethylenglycol)

n

O

O

H2C–CH–CH2–O –CH2–CH2 –O –CH2 –CH2 –O–CH2 –CH –CH2 n

OH

OH

OH

–O–Si–(CH2)3–NH –CH2–CH–CH2–O–CH2–CH2–O–CH2–CH2–O–CH2–CH–CH2–NH– n

E

OH

Fig. 4. Introduction of oxirane groups to poly(ethylenglycol) (PEG) by activation with epichlorohydrin. The activated polymer (ECH-PEG) can be used as coupling agent between enzyme and glass support.

3. Spacer-modified glass can be dried by air; even though further use in wet state is possible, it would hamper the mass balance due to the residual water. Storage under refrigeration is recommended. 3.2.2. Modification with PEG and PEG-BSA-PEG

The spacer length is important for the regulation of the enzyme and cofactor activity, especially if mesoporous materials with different pore diameters are going to be used. Polymers like poly(ethylenglycol) (PEG) of any length can be used as spacer after being activated with ECH (epichlorhydrin) (15). The degree of activation with ECH can be monitored by pH measurement. ECH introduces a highly reactive oxirane group to both ends of the polymer chain. ECH-activated PEG can easily be attached to the activated glass before the enzyme is bound via the second oxirane group (Fig.  4). Previous studies have shown that the adjustment of the flexibility of the spacer also plays a role in the regulation of the catalytic activity of a multiple enzyme system. Therefore, the utilization of bovine serum albumin (BSA) as additional spacer might be beneficial. This can be achieved after the activated glass is treated with ECH-activated PEG, followed by coupling with BSA that is subsequently treated with ECH-activated PEG (in such case, the spacer is comprised of PEG–BSA–PEG) before the enzymes and cofactor are finally attached. 1. One gram (1 g) of PEG (10,000 Da) is dissolved in 10 mL epichlorohydrin (ECH) and 10  mL water. The resulting mixture is vigorously stirred for 12 h at room temperature. 2. On completion of the reaction, the ECH-activated PEG is precipitated by evaporation of the solvent through nitrogen blowing. By this process, excessive ECH and residual water

Nanoporous Silica Glass for the Immobilization of Interactive Enzyme Systems

43

are removed. The dry epoxy-terminated PEG can be stored at 4°C. 3. One gram (1  g) of activated glass is suspended in 8  mL of solution D containing 1 g of solubilized ECH-modified PEG and incubated overnight at room temperature under gentle stirring (see Notes 6 and 7). 4. Activated and spacer-modified glass is removed from the solution by centrifugation and at least three times washed with water to separate excessive ECH-modified PEG. The glass particles should be dried and kept at 4°C prior to further use. Steps 5–7 are optional. 5. One gram (1 g) of activated and spacer-modified glass is suspended in 8 mL of solution E and incubated for 24 h while gently shaken at room temperature. 6. The BSA-attached glass is recovered by centrifugation and washed with water until no more protein can be detected in the supernatant (using the Bradford assay). 7. Repeat steps 3 and 4. 3.3. Attachment of Enzymes and Cofactor to Glass

Attachment requires the contacting of activated glass support and an aqueous solution containing the enzyme and/or the cofactor. Based on our studies (7), a two-step approach comprising a first contact with the cofactor and then the contact between enzyme and glass support comes up in the highest catalytic efficiency since it best guarantees that enough cofactor is loaded. Depending on the amount of enzyme and the nature of the spacer used, typically, final loadings are in the range of 1–2 mg NADH and 0.1–0.2 mg enzyme per gram of support (typically immobilization degree equals to 15–30%). 1. One gram (1 g) of the activated glass material is mixed with 4 mL of solution F. The mixture is gently shaken for 48 h at room temperature (see Notes 5 and 6). 2. NADH-attached glass is removed from the solution by centrifugation and washed at least three times with cofactorfree buffer (solution D) to separate excess NADH. By measurement of the UV absorbance at 340 nm, the attachment and washing can be monitored. Based on the mass balance of the measured concentration, the amount of immobilized NADH can be calculated. If storage of NADH-attached glass support is necessary, we recommend refrigeration to 4°C, but storage period should not exceed a couple of days. 3. For the immobilization of the enzyme, 1 g NADH-attached glass is contacted with 4 mL of the enzyme containing solution G (see Note 4). The mixture is kept for 24 h at 4°C while it is gently shaken.

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4. The enzyme-NADH-attached glass support is removed from the solution by centrifugation, followed by resuspension in 4-mL enzyme-free buffer (solution D) and centrifugation for at least three times to wash off excess unbound enzyme. In this context, it is recommended to analyze the protein content in the washing water of each step to make sure that all nonimmobilized enzyme is removed (see Note 7). Additionally, the mass balance can be used to calculate the enzyme loading and immobilization degree (amount of enzyme immobilized onto the support compared to the total protein applied). Due to the occasional delicate stability of wet catalyst systems, it is recommended to use the immobilizates promptly. If necessary, storage in the refrigerator at 4°C for a maximum of a couple of days is possible. 3.4. Specific Activities of Immobilized and Free Enzyme 3.4.1. Activity Assay

In order to judge the efficiency of the immobilized biocatalysts especially compared to their free nonimmobilized counterparts, it is necessary to use a reliable quantitative assay for the determination of the catalytic activity and the protein concentration (see also Note 7). It is necessary to distinguish between the specific activity (U/mg protein) of the native enzyme, the specific activity of the immobilized protein, and the apparent activity of the immobilizate (U/mg enzyme-attached glass). A prerequisite for the denomination of those activities is the knowledge of the protein amount present in the enzyme solution before and after being exposed to the activated glass, as well as the protein amount in the washing water (which enables the calculation of the enzyme amount immobilized onto the mesoporous glass (enzyme loading) and the immobilization degree (fraction of total protein getting immobilized)). Typically, when PEG was used as spacer, the reaction rate for pyruvate ranged from 0.6 to 1.2  mmol/L/h/mg enzyme and from 0.25 to 0.50 mmol/L/h/mg enzyme for glucose. The NADH turnover number (mmolsubstrate/L/h/mmolNADH) is defined as the normalized reaction rate with respect to the amount of cofactor and ranged from 40 to 100 1/L/h NADH for pyruvate and from 15 to 40 1/L/h for glucose. When glutardialdehyde was used as spacer, the observed values for reaction rate and turnover number were, on average, one order of magnitude smaller (see Note 8). 1. The substrate solution H used for the determination of the LDH and GDH activity comprises 1 mM pyruvate and 1 mM glucose solubilized in 50 mM phosphate buffer (solution D). An amount of 250 mg of the immobilized glass particles is suspended in 5 mL of the substrate solution and vigorously stirred (see Note 5). 2. Samples of 0.5  mL are withdrawn periodically and filtered using 0.2 mm PTFE syringe filters. The filtrate undergoes a

Nanoporous Silica Glass for the Immobilization of Interactive Enzyme Systems

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HPLC analysis for the determination of pyruvate concentration, whereas the glucose concentration is determined by using an enzyme-based assay (peroxidase and glucose oxidase). Time course of the measured concentrations is used to evaluate the initial reaction rate. It is recommended to conduct the experiment at least in triplicate to obtain reliable results. 3.4.2. Bradford Protein Assay

The Bradford assay utilizes a colorimetric reagent for the detection and quantification of total protein (16). Protein in solution binds to the reagent (Coomassie Brilliant Blue G250) accompanied by a spectral shift from 465 to 595 nm. The spectral shift is proportional to the amount of protein present in solution. Sample protein concentration is determined by calibration with BSA as standard protein. The applied calibration range is between 0 and 20  mg/mL. The sample solution was diluted to an appropriate concentration and 500  mL were added to a 1.5-mL cuvette followed by the addition of 500 mmL Bradford reagent.

3.4.3. Analysis of Substrates and Products

For the analysis of all involved substrates and products, we propose as a method of choice to use an HPLC system equipped with an UV and RI detector. Thereby, for the column, a stationary phase based on a polyvinyl alcohol-based gel with incorporated amino groups with a polar eluent (mixtures of water, acetonitrile, and ethanol) is advised (17). Admittedly, this configuration is not a common one, which is why a method that can be conducted in most laboratories is described below. 1. For the detection of pyruvate and lactate, an HPLC system should be equipped with a UV-VIS detector (wavelength adjusted to 215 nm) and a C-18 column (4.6 mm × 250 mm). 2. As isocratic mobile phase, a 50/50 (v/v) mixture of acetonitrile and water can be used at a flow rate of 1  mL/ min. It is important to lower the pH of the eluent to about pH 2.0 by adding an inorganic acid (e.g., sulfuric acid) for the complete protonation of the organic acid. 3. Calibration is done with standards of known concentrations (injection volume 10 mL). Samples themselves are diluted to appropriate concentrations compliable with the available HPLC system. All analyses should be conducted in triplicates to minimize standard errors. As the above-mentioned HPLC method does not allow the detection of glucose, its concentration in the reaction mixture can be determined by its enzymatic oxidation with glucose oxidase (18). This reaction is accompanied by the release of H2O2, which is subsequently decomposed by peroxidase under the concomitant oxidation of the colorant o-dianisidine (turns from colorless to brown).

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1. For the calibration, different standards of known glucose concentration in the range of 0–1 mM are prepared. 2. A sample volume of 25 mL sample is added to 0.5-mL water, which is subsequently added to 5 mL of the reactant solution (solution I). The reactant solution comprises both enzymes, the glucose oxidase (5,000 U/mL GOx) and the peroxidase (1,000 U/mL PO), as well as o-dianisidine dihydrochloride (40 mg/mL). 3. The final mixture is mixed while avoiding light exposure for a total time of 30  min. Finally, the absorbance at 450  nm is measured and, based on the calibration, the sample concentration is calculated. Each sample should be analyzed in triplicate to minimize the standard errors.

4. Notes 1. The protocol given above can easily be applied to other enzymes. The only restriction might be given if the enzyme activity is tremendously lowered by covalent attachment via aldehyde or oxirane groups. If literature does not provide information about that, a simple pretesting should be carried out to verify the putative toxicity of the coupling agent. 2. The handling of mesoporous glass particles can be awkward owing to their small particle size. Therefore, the removal from solution can be best achieved by moderate centrifugation as filtration is typically accompanied by a higher loss of material. For convenience, 15-mL centrifuge tubes are recommended for all protocol steps dealing with the glass support. The centrifugation can be done at a moderate speed of 4,500 rpm for 5 min. 3. As for every multistep protocol, it has to be considered that each step contributes to the overall standard deviation (Fig. 5). Therefore, it is recommended that at least three different batches of enzyme-NADH-attached glass particles be prepared. Otherwise, it might be difficult to judge the potential of the method. 4. Volume of solutions and amounts given in this protocol are not mandatory. Volume is less a critical issue as long as the support material is completely covered. The optimum concentrations cannot be defined bindingly and have to be evaluated empirically. For example, the best enzyme protein concentration depends on the purity of the formulation used, what only can be determined experimentally.

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Mesoporous silica glass Activation with APTMS

Attachment of GDA

Activation of PEG with ECH

Attachment of ECH-PEG

Cofactor attachment

Cofactor attachment

Enzyme attachment

Enzyme attachment

BSA attachment Attachment of ECH-PEG

Cofactor attachment

Enzyme attachment

Fig. 5. Flow scheme summarizing the crucial protocol steps for nanoscale immobilization.

5. As inorganic particles are brittle materials, the use of a stirring bar might “grind” the particles. Therefore, better results can be obtained if the glass particles are suspended by gently shaking the reaction tubes. 6. Occasionally, the inorganic particles show a tendency to clump, and the resuspension of dried particles seems to be difficult. The best remedy for this is ultrasonication for 1 min. 7. If the results are poorly reproducible or misleading, one probable reason is the leaching of nonimmobilized enzyme that was not covalently attached to the carrier and also not totally removed by washing. To verify this, we recommend assessment of the soluble activity of the supernatant by withdrawal and subsequent incubation prior to analysis. In such case, even if no particles are present, substrate consumption will be observed. 8. If results are unsatisfying, one possible reason might be impurity of the glass support. If the purity is not certified by the supplier of the mesoporous glass, it is recommended that the support material be cleaned by heating to a temperature high enough to burn off any organics adsorbed. In such case, it might be necessary to rehydrate the surface prior to further use.

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References 1. Ansorge-Schumacher M.B. (2008) Immobili­ zation of biological catalysts. Handbook of Heterogeneous Catalysis (2nd Edition) 1, 644–55. Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim. 2. Hanefeld U., Gardossi L., Magner E. (2009) Understanding enzyme immobilization. Chem Soc Rev 38, 453–68. 3. Reetz M.T., Tielmann P., Wiesenhoefer W., Koenen W., Zonta A. (2003) Second generation sol-gel encapsulated lipases: robust heterogeneous biocatalysts. Adv Synth Catal 345, 717–28. 4. Wang P. (2009) Multi-scale features in recent development of enzymic biocatalyst systems. Appl Biochem Biotechnol 152, 343–52. 5. Wang P. (2006) Nanoscale biocatalyst systems. Curr Opin Biotechnol 17, 574–9. 6. Kim J., Grate J.W., Wang P. (2008) Nano­ biocatalysis and its potential applications. Trends Biotechnol 26, 639–46. 7. El-Zahab B., Jia H., Wang P. (2004) Enabling multienzyme biocatalysis using nanoporous materials. Biotechnol Bioeng 87, 178–83. 8. Wang P., Dai S., Waezsada S.D., Tsao A.Y., Davison B.H. (2001) Enzyme stabilization by covalent binding in nanoporous sol-gel glass for nonaqueous biocatalysis. Biotechnol Bioeng 74, 249–55. 9. Yi Y., Kermasha S., Neufeld R. (2008) Nanoporous sol-gel supports enzymatic hydrolysis of chlorophyll in organic media. ACS Symp Ser 986, 199–213. 10. Weetall H. (1993) The activation of inorganic carriers by silanization. Biosens Bioelectron 8, x–xi.

11. Liu W., Zhang S., Wang P. (2009) Nanoparticle-supported multi-enzyme biocatalysis with in situ cofactor regeneration. J Biotechnol 139, 102–7. 12. Liu W., Wang P. (2007) Cofactor regeneration for sustainable enzymatic biosynthesis. Biotechnol Adv 25, 369–84. 13. El-Zahab B., Gonzalez D., Wang P. (2004) Dendrimer-supported multienzymatic biocatalysts with in situ cofactor regeneration. Abstracts of Papers, 228th ACS National Meeting, PMSE-037, Philadelphia, PA. 14. Aldercreutz P. (1996) Cofactor regeneration in biocatalysis in organic media. Biocatalysis Biotransformation 14, 1–30. 15. Andrews B.A., Head D.M., Dunthorne P., Asenjo J.A. (1990) PEG activation and ligand binding for the affinity partitioning of proteins in aqueous two-phase systems. Biotechnol Tech 4, 49–4. 16. Bradford M.M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of proteindye binding. Anal Biochem 72, 248–54. 17. Roda A., Sabatini L., Barbieri A., Guardigli M., Locatelli M., Violante F.S., Rovati L.C., Persiani S. (2006) Development and validation of a sensitive HPLC-ESI-MS/MS method for the direct determination of glucosamine in human plasma. J Chromatogr , B: Analyt Technol Biomed Life Sci 844, 119–26. 18. Kapustka L.A., Annala A.E., Swanson W.C. (1981) The peroxidase-glucose oxidase system: a new method to determine glucose liberated by carbohydrate degrading soil enzymes. Plant Soil 63, 487–90.

Chapter 6 Enzyme Stabilization and Immobilization by Sol-Gel Entrapment Allan E. David, Arthur J. Yang, and Nam Sun Wang Abstract While biocatalysts show tremendous potential for the industrial production of fine chemicals, their integration into large-scale processes has been slow. One of the main reasons for slow acceptance in industry is the inherent instability of the enzymes. Recent developments in bioengineering have shed some light on methods of improving enzyme stability. One method that has been used for many decades, successfully to varying degrees, has been the immobilization of enzymes. To this regards, silica gels have attracted much attention because of the ease of surface functionalization, high surface areas, mechanical and thermal stability, and resistance to both chemical and biological attack. We have previously shown the immobilization of invertase on silica gels with high immobilized activity and significantly improved stability. Here, we provide greater details on the methods for effecting the immobilization. Key words: Enzyme immobilization, Entrapment, a-Amylase, Silica gel, Sol-gel, Silicic acid, Aminopropyltriethoxy-silane, Glutaraldehyde, Cross-linker

1. Introduction Biocatalysts have drawn considerable attention for use in industrial process because of their high chemical precision, compared to organic chemistry. Recent advancements have made the use of enzymes in industrial processes much more feasible (1). Developments in bioengineering have allowed for the overexpression of selected enzymes in fermentations and, coupled with improved purification techniques, lowered the cost of enzymes (2, 3). In addition, the developments in protein engineering and advanced screening techniques have introduced a large variety of designed enzymes providing greater selection for the improvement of particular reactions (4). Most recently, the development of nonaqueous enzymatic processes has drawn interests from many groups due to Shelley D. Minteer (ed.), Enzyme Stabilization and Immobilization: Methods and Protocols, Methods in Molecular Biology, vol. 679, DOI 10.1007/978-1-60761-895-9_6, © Springer Science+Business Media, LLC 2011

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improved product and enzyme recovery (5–7). Despite the fact that over 3,000 enzymes have been isolated and characterized, only a few are currently utilized in industrial processes. While the potential for enzymes to catalyze commercially important reactions is enormous, integration into industrial processes is limited by enzyme instability. In general, enzymes are easily denatured by changes in temperature or pH, and susceptible to attack by both chemical and biological agents. Methods to overcome these limitations include protein engineering, in which amino acid sequences are modified in order to obtain a more stable enzyme, and the isolation of new enzymes from extremophiles – organisms living in extreme conditions (8). Other methods to improve enzyme stability include chemical modification of the enzyme surface or the immobilization of enzymes to a solid surface. We have previously demonstrated that chemically surface-modified silica gels (CSMG) can serve as an ideal matrix for industrial enzyme immobilization, retaining a high degree of enzyme activity (9). In this chapter, we will focus on the immobilization of enzymes in silica gel matrices. Enzyme immobilization involves certain costs associated with the cost of materials (e.g., polymer matrix and cross-linkers) and processing time. The two major driving concerns in an industrial scale process are to lower the unit cost and to increase the unit production per fixed time. Immobilization becomes economically feasible only if it allows the repetitive use of enzyme in multiple batches to reduce the unit production cost. Further, immobilized enzyme beads can be packed into a bed or a column for continuous flow-through reactions, if applicable, to reduce costs associated with otherwise labor-intensive processes. Such a reactor also simplifies the separation of enzyme from product: alleviating some downstreamprocessing costs. The appropriate immobilization matrix is chosen based on several different properties which affect the production process (10): ●●

●●

●●

Surface area and porosity: it is desirable to have materials with high surface areas (>100  m2/g), for high enzyme loadings, and high porosity to provide enzyme access for the substrate. Pore sizes >30  nm are ideal for the diffusion of enzymes during the immobilization process. Surface functional groups: the degree of enzyme loading onto a carrier matrix also depends on the loading density of functional groups on the surface and its distribution. Choice of functional groups also affects the activity yield and material stability. Mechanical and chemical stability: many immobilized enzyme operations are conducted as a stirred-tank or packed-bed reactor. To prevent enzyme loss, the matrix integrity must be

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maintained under the shear-stresses or back-pressures present in these reactors. In addition, the matrix must be resistant to chemical degradations which will also result in the loss of enzyme. ●●

●●

●●

Size and shape: for simplifying handling of the immobilized enzyme (i.e., stirring, filtration), it is ideal to have particles of uniform shape and size. For this reason, a uniform spherical matrix is preferred and also results in the reduction of back-pressures in column reactors. In addition, spherical particles are also more easily characterized for modeling purposes. Microbial resistance: a major concern of any immobilized enzyme process is the presence of microbes. The durability of the carrier is often determined by its resistance to microbial degradation. Hydrophobic/hydrophilic nature: the compatibility of the support with the liquid phase is important to insure the free exchange of substrate and product between the matrix and bulk phase. It can also determine the life-time of the matrix due to the surface adsorption of materials through nonspecific interactions.

Immobilization allows the enzyme to be used in a continuousflow mode which provides several benefits: (1) continuous removal of products from the reactor, which can be beneficial for systems that suffer from product inhibition; (2) product separation through column retention, reducing the burdens for downstream purification by affinity columns; and (3) increased enzyme stability will allow the enzyme to maintain activity for longer periods of use and in more extreme conditions. 1.1. Methods of Enzyme Immobilization

Enzyme immobilization, the restriction of mobility, can be accomplished in a variety of ways, including entrapment, crosslinking, or physical attachment (11). Enzyme entrapment is accomplished through the polymerization of a matrix from a solution containing both the monomer and enzyme. As shown in Fig. 1a, entrapment results in the physical entrapment of enzymes within a cage-like matrix. While, in general, enzyme entrapment does not involve any modification of the enzyme and therefore avoids adversely affecting activity in this way, denaturing of the enzyme may occur due to its exposure to matrix precursors or due to changes in microenvironment during matrix polymerization. We will provide an example for the entrapment of an enzyme in silica gels later in this chapter. It should be noted that this method, enzyme entrapment, can be susceptible to leaching of enzymes out of the matrix by diffusion when pore diameters are larger than the enzyme.

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Fig. 1. Methods of enzyme immobilization include (a) entrapment, (b) cross-linking, and physical attachment by (c) adsorption, (d) ionic interaction, and (e) covalent binding.

Cross-linking, which involves the attachment of enzymes to each other (Fig. 1b), is another method for immobilizing enzymes (12). However, in many cases, the extensive interactions required, whether electrostatic, chemical, or physical in nature, tend to significantly reduce enzyme activity by hindering formation of the enzyme’s optimum folding conformation. In addition, because the material properties are governed by the enzyme, which is the major component in the cross-linked beads, mechanical strength can be fairly low: limiting the cross-linked enzymes to low system pressure reactors. The physical attachment of enzymes to a solid matrix, by adsorption, ionic, or covalent interaction (see Fig. 1c–e, respectively), provides several advantages compared to entrapment and cross-linking. Due to the stronger interactions with the matrix, physical attachment provides greater material stability against leaching when compared to entrapment. However, if the immobilization is accomplished by adsorption or ionic interactions, leaching may still occur depending on system conditions (i.e., pH, temperature, and solution ionic strength). The covalent attachment of enzymes to the matrix is preferred for some applications because of its inherent stability. Covalent immobilization is accomplished through the binding of reactive groups on the enzyme with a chemically active surface. Covalently immobilized enzymes are suitable in a variety of reaction environments with no significant increase in leaching due to the strong binding. However, the strong binding may also inhibit enzyme movement and thus reduce its activity by preventing conformational changes, which can also be the means of enzyme stabilization.

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Enzymes are composed from a selection of 20 different amino acids with the majority of these acids having relatively inert side chains composed of hydrocarbons. However, as seen in Fig.  2, nine of the amino acids have chemically active functional groups (i.e., amino, thiol, hydroxyl) on their side chains. The configuration of an enzyme is determined by the interaction of these side groups among themselves and with the environment (e.g., disulfide binding, amide bonds, and hydrophobic/hydrophilic interactions). It is also through these interactions that enzymes can be attached to a solid surface. Although the thiol group of cysteine is the more potent nucleophile, the amino groups are the more important target because of their relative abundance in proteins. In addition, sulfhydryl groups are generally found in pairs that form disulfide bonds and strongly influence protein structure. Modification of these sulfide bonds may have detrimental effects on enzyme activity. Therefore, the preferred binding site is the amino residue on lysine side chains (10). Many enzyme immobilization techniques involve SN2-type, nucleophilic substitution reactions. A typical enzyme immobilization system can consist of four different parts: the matrix; a surface modifying agent, if the matrix does not possess reactive functional groups; a cross-linking agent, which attaches the enzyme to the matrix; and the enzyme itself. An ideal situation would allow immobilization to occur by simply mixing all matrix

Fig. 2. Amino acids with chemically active side chains that can be utilized to covalently immobilize enzymes.

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and reactive precursors together with the enzyme. Polymerization of the matrix would then allow for enzyme immobilization at high loadings and even distribution. This is not possible in the vast majority of cases because nonspecific interactions between the reactive groups can result in the formation of undesirable bonds. Here, we will discuss the utilization of sol-gel silica chemistry for the immobilization of enzymes. A sol, by definition, is a suspension of solid particles, with size between 1 and 1,000 nm, in a liquid continuous phase. The sol-gel process refers to the creation of a continuous, solid network through a change of interactions between the colloidal particles; changing the systems characteristics from that of a liquid to that of a gel. The result is a bi-continuous system composed of the continuous, interpenetrating solid and liquid phases. Freshly prepared silicic acid is composed of silica particles with very small particle size ( 4. 6. The silica sol has limited stability, due to cocondensation between particles resulting in the formation of a gel, and the solution must therefore be freshly produced for enzyme immobilization (see Note 5).

3.2. Entrapment of the Enzyme a-Amylase in Silica Sol-Gel

Entrapment of enzymes in a matrix requires that enzyme activity be preserved in the presence of all precursors and conditions used for the immobilization. In the case of entrapment in silica sol-gels, since the silicic acid transitions from acidic state (pH 3) to neutral during gelation (as detailed below), it is important to insure that the changing pH does not cause enzyme denaturization.

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For example, when the enzyme a-amylase was mixed directly with silicic acid and the gel formed, no retained enzymatic activity was observed. While the following procedure illustrates a method for retaining the activity of a-amylase entrapped in silica sol-gels, it can also be applied to other enzymes. 1. a-Amylase is added to 100 mM acetate buffer (10 mL, pH 7) at a final concentration of 1  mg/mL and set aside. The enzyme concentration can be increased or decreased to vary the entrapped enzyme loading. 2. Silicic acid (100 mL of 10 wt%, pH ~ 3) is poured into a beaker and the solution magnetically stirred. 3. A pH probe is placed in the solution to allow continuous monitoring of the system. 4. Since the gelation of silica sol is on the order of minutes at 4 100 U SOD, and less significant effects at lower doses. Neither SOD (25–200 U) nor PEG–SOD (100 U) in solution demonstrated the neuroprotective effect under similar conditions. The neuroprotective effect of SOD-nanoparticles was seen up to 6 h after H2O2-induced oxidative stress, but the effect diminished thereafter. The suggested mechanism of efficacy of SODnanoparticles appears to be due to the stability of the encapsulated enzyme and its better neuronal uptake after encapsulation. More recent in vivo study by the same group (58) evaluated neuroprotective efficacy of SOD-loaded PLGA nanoparticles in a rat focal cerebral ischemia-reperfusion injury model. The authors studied three routes of administration via intravenous (tail vein), intrajugular, and internal carotid arterial routes. They found that ~0.1% of nanoparticles were uptaken by the brain in the case of intravenous and intrajugular administration, and ~1.5% of nanoparticles were uptaken by the brain in the case of administration via the carotid artery. The nanoparticles administered via the carotid arterial route improved the survival rate (75% vs. 0% in controls) and showed good neuroprotective efficacy in this in vivo model. It appears that improved delivery to the brain was critical for in vivo efficacy. However, intracarotid arterial route may not be a preferable administration route and is unlikely to be beneficial if CNS injury is not located in brain (e.g., in the case of secondary spinal cord injury). Thus, there is an apparent need for the development of more efficient ways of delivery of neuroprotective enzymes to CNS. One of the significant achievements was the demonstration that functionalized polybutylcyanoacrylate (PBCA) nanoparticles can penetrate through blood–brain barrier and be used for drug and gene delivery to central nervous system (CNS) (59, 60). Reukov et  al. 61) showed that SOD as a free radical scavenger and NR1 antibody for targeting to the neurons can be simultaneously attached to the PBCA nanoparticles without considerable changes in enzymatic activity or receptor-binding ability.

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Neuroprotective action of anti-NR1-SOD-nanoparticle conjugates was studied using rat neuron cell culture. Rat DRG neurons were challenged by superoxide (generated using Xanthine/Xanthine oxidase system) in the presence and the absence of the conjugates. Viability of the cells has been characterized using live/ dead assay. Results are indicative of strong neuroprotective action of the conjugates.

4. Targeted Delivery of Thrombolytic Enzymes

4.1. Targeted Delivery Using Magnetic Carriers

Currently, recombinant tPA is used clinically in postmyocardial infarction treatment to inhibit blood clot formation. Unfortunately, tPA also acts upon plasminogen in circulation generating systemic plasmin and rendering the treated patient highly vulnerable to hemorrhage (62, 63). Indeed, administering recombinant tPA is associated with a high patient mortality rate (6.3% for 30-day mortality) including fatal cerebral hemorrhage in 1.5% of patients (64). While beneficial effects of tPA-based thrombolytic therapy have already been established, a decrease in the associated mortality rate would vastly improve therapeutic outcomes. This requires a mechanism for direct delivery of tPA to the clot site, where it can act solely upon fibrin-bound plasminogen and minimize systemic plasminemia. Targeted delivery of thrombolytic enzymes, including tPA, streptokinase or urokinase-type Plasminogen Activator (uPA), has been extensively studied to improve outcome of cardiovascular disease. A number of attempts have been made to enhance targeted delivery of these enzymes to the blood clot sites. These methods are (a) direct application of the drug to the affected site using a catheter, (b) antibody-based targeting to the blood clot components, (c) targeting via proteins other than antibodies, (d) targeting using magnetic carriers, (e) liposome-based targeting, (f) targeting using polymeric carriers, and (g) targeting using red blood cells as carriers. Several groups (65–69) focused on the use of magnetic carriers with loaded drug for targeted delivery to the vicinity of a blood clot. Authors of ref. (65) used red blood cells preloaded with a magnetite-based ferromagnetic colloid and aspirin. These cells were injected and moved to the proximity of a damaged blood vessel using an external SmCo magnet. It was shown that this approach can be used to inhibit or completely prevent clot formation. Torchilin et al. (67) studied streptokinase covalently attached to dextran-coated magnetic carriers in a dog carotid artery thrombosis model. They found that normal blood flow in the arteries

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can be restored 5 h post surgery if a strong SmCo magnet was placed in the vicinity of the animal’s carotid artery. Efficacy of magnetic carriers of thrombolytic agents has also been demonstrated in vivo in rabbit (68) and rat (69) models. 4.2. Targeted Delivery Using Liposomes

Liposomes – artificial mono- or multilamellar phospholipid vesicles of various size and composition – have been recognized as pharmaceutical carriers with great practical potential (70, 71). Liposomes obtained from natural phospholipids are completely biocompatible and biodegradable. They can be loaded with a water-soluble drug into their aqueous compartment or with a lipid-soluble drug in the membrane providing isolation from the external environment and preventing undesirable side effects. McPherson et al. (72) studied liposomes loaded with tPA in an in-vitro thrombolysis model. tPA was loaded in the aqueous compartment of the liposomes and was released upon sonication. It was found that thrombolytic activity of the enzyme released from the liposomes was higher than that of free tPA and can be further increased with additional ultrasonic treatment. The reason for this enhanced thrombolytic activity is probably elimination of diffusion limitations upon sonication, rather than effect of liposome carriers on tPA function. However, it would be interesting to see results of in  vivo studies of tPA-loaded liposomes since use of these carriers can be expected to considerably prolong half-life of tPA in the circulation and the ability to achieve tPA release at a desired time and location via sonication could be an attractive feature for therapeutic applications.

4.3. Targeted Delivery Using Polymeric Nanoparticles

A novel approach for targeted delivery of thrombolytic enzymes is based on the use of polymeric nanoparticles as carriers. In this case, the enzyme and the targeting ligand can be simultaneously attached to the surface of nanoparticles. Our group reported simultaneous attachment of tPA and antifibrin antibody to polystyrene latex nanoparticles (73). In vitro studies of fibrinolysis showed that tPA-antifibrin–nanoparticle conjugates were only slightly less potent than free tPA. Interestingly, tPA activity attached to nanoparticles in the absence of clots was approximately fourfold lower than that of the free enzyme (Fig. 3). The reason for the observed effect is probably enhanced activation of tPA by fibrin when antibody-directed binding brings it into close proximity with a clot. This property can be of importance for therapeutic applications because nanoparticles can be expected to dissolve clots at approximately the same rate as free tPA while simultaneously cleaving much less plasminogen in the circulation, hence lowering the risk of systemic toxicity. This system could therefore become a promising agent for thrombolytic treatment.

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a

100-100 50-100 100-50 tPA

Absorbance at 405 nm

3

2

1

0 0

200

400

600

Time2

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1000

(sec)2

b

Fig. 3. (a) Activity of tPA conjugated to latex nanoparticles compared to that of free tPA. (b) In vitro fibrinolytic activity of samples 100:100, 50:100 and 100:50 compared to that of free tPA (n = 4 for each point).

5. Antibacterial Enzymes and Peptides

Increased antibiotic resistance in microorganisms has become a major problem in clinical practice, which heavily relies on the use of antibiotics for treatment of infectious diseases (74). Increased resistance of a number of human pathogens, including M. tuberculosis,

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E. faecium, S. aureus, and P. aeruginosa to clinically important antibiotics has caused a crisis in the treatment and management of a number of bacterial infections. The decreased effectiveness of current antibiotic treatments due to resistance has spurred the search for alternative approaches to antimicrobial therapy. Increasingly, research is aimed towards the use of natural antimicrobial enzymes and peptides as a potential solution to the resistance problem (75). Naturally occurring antimicrobial enzymes and peptides (AMPs) have attracted much attention during the past two decades because of their high level of activity, broad antimicrobial spectrum, and the low rate of development of resistance to these agents by bacteria. Antibacterial peptides are efficient against many multidrug-resistant pathogens and, therefore, can serve as potent alternatives to conventional antibiotics. Several antimicrobial peptides are currently being tested in clinical trials. Thus, these nature-inspired antimicrobial agents are promising candidates for the development of novel therapeutic agents for treatment of infectious diseases (76). Overall, great enthusiasm exists regarding the prospect of developing AMPs as a new generation of medications for treatment of a variety of infections. In spite of this high promise, use of AMPs in the clinical setting is associated with several potential drawbacks. Because of their peptidic nature, AMPs could present the following problems: (1) high manufacturing costs, (2) low stability and short half-life, (3) potential toxicity and unwanted systemic reactions (aggregation, immunoreactivity), that may arise when attempts are made to use those peptides as antibacterial agents (23). The use of antimicrobial enzymes covalently attached to nanoparticles can address the latter two problems because of enhanced stability of protein–nanoparticle conjugates and the possibility of targeted delivery. 5.1. Nanoparticles As Carriers of Antibacterial Enzymes and Peptides

A number of polymeric nanoparticles have been employed as antibiotic carriers. Among these were polyacrylamide, polyalkylcyanoacrylate, and polylactide-co-glycolide nanoparticles loaded with such antibiotics as ampicillin, ciprofloxacin, and rifampicin (77–80). In the case of low-molecular-weight antibiotics, the role of the carriers is usually restricted to sustained release of the drug. Antibacterial activity of nisin-loaded polylactide nanoparticles against Lactobacillus delbrueckii has also been evaluated (81). In most of these cases, the role of nanoparticles was to provide a sustained release of the loaded drug; however, targeted delivery of antibiotics to infected macrophages and to liver has been utilized using nanoparticles and liposomes (76, 77). Such targeting was possible because of the increased uptake of polymeric nanoparticles by reticuloendothelial system.

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Recently, specific targeted delivery of polymeric nanoparticles either unloaded or loaded by an antimicrobial drug, to bacteria have been studied. Mannosylated or galactosylated polystyrene latex nanoparticles were shown to result in agglutination of mutant strains of E. coli overexpressing mannose and galactose surface receptors, respectively (82–84). These studies have been thus far restricted to the model E. coli mutants; however, the authors envision development of nanoparticles coated by oligosaccharides that will specifically recognize desired bacterial strains and kill them by sticking to bacterial cell walls and causing bacterial agglutination. This approach is currently at the very early stage of development, and its prospective clinical applications are unclear. Potential problems could be the need for high doses to achieve agglutination of bacteria and impossibility to treat colonized bacteria. Effect of antibody-directed targeting to Gram-positive Listeria monocytogenes has recently been studied by the same group for lysozyme attached to polystyrene latex nanoparticles simultaneously with anti- L. monocytogenes antibody (85). The authors observed significantly (p 70-nm Au), the Au surface should look planar to the protein. (b) Transmission electron micrograph of Au~cyt.c superstructure within Au~cyt.c-SiO2 composite aerogel – the silica domains of the aerogel, the protein superstructure, and the nucleating gold colloid are distinguishable (Reproduced from ref. 13 with permission from the American Chemical Society, copyright 2003.).

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within silica during the sol-to-gel transition, and processed to form aerogels in which cyt. c retains its characteristic visible absorption (12–15). The Au~cyt. c superstructures even exhibit rapid gas-phase recognition of nitric oxide (NO) within the aerogel matrix, facilitated by the high-quality pore structure of the aerogel (13). These bioaerogels do not need to be kept wet or stored at 4°C to maintain protein viability. The organization of protein within the Au~cyt. c superstructure imparts vital protection against protein denaturation during the harsh physical and chemical processing conditions necessary to form the aerogel. Thousands of cyt.c molecules per gold nanoparticle are stabilized within each superstructure. Prior work using guanidinium hydrochloride to unfold the protein demonstrated that even in the absence of the aerogel scaffold, the protein is stabilized to unfolding relative to cyt. c in buffered solution, providing further evidence of the existence of the Au~cyt. c superstructure in open medium (13). Such stability makes these superstructures interesting both from a fundamental standpoint and from an applied standpoint because there is potential to make use of biomolecular composite aerogels for bioanalytical applications.

2. Materials 2.1. Silica Sol Preparation

1. Polypropylene disposable beakers (50 mL, Fisher). 2. Methanol (GC Resolv grade) (see Note 1). 3. Tetramethoxysilane (TMOS; also referred to as tetramethylorthosilicate (98%, Aldrich)). 4. Ammonium hydroxide solution (ACS reagent, 28.0–30.0% NH3 basis, Sigma-Aldrich). 5. General-purpose polypropylene scintillation vials (16  mm ×  57  mm, volume size 6.5  mL, Sigma Aldrich) with bottom end sliced off. 6. Generic plastic wrap. 7. Parafilm M™ laboratory wrapping film (Fisher).

2.2. Au~cyt. c Superstructure Preparation

1. Sodium phosphate, Monobasic, Reagent ACS. 2. Sodium phosphate, Dibasic, Anhydrous Powder. 3. Cytochrome c (cyt. c) from equine heart (≥95% based on Mol. Wt. 12,384, Sigma Aldrich) used as received and stored at −20°C. All buffered solutions prepared are stored at 4°C (see Note 2). 4. Colloidal gold sol, 5 or 10 nm (20 mL, stored in the dark at 4°C, BBInternational) (see Note 3).

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5. Disposable cuvette or quartz cuvette (1-cm pathlength). 6. Glass 20-mL Scintillation Vials (O.D. × Height (with cap): 28 mm × 61 mm, Wheaton). 2.3. Au~cyt. c Silica Sol–Gel Preparation

1. Polypropylene disposable beakers (50 mL, Fisher). 2. Ethyl alcohol, absolute, 200 proof, 99.5% A.C.S. reagent. 3. Acetone, HPLC grade. 4. Plastic syringe plunger. 5. Glass 20-mL Scintillation Vials (O.D. × Height (with cap): 28 mm × 61 mm, Wheaton).

2.4. Au~cyt. c Silica Aerogel Preparation

1. Carbon dioxide, siphon tube cylinder (Tech Air).

2.5. UV–Visible Spectroscopy of Au~cyt. c Silica Aerogels

1. Au~cyt. c silica aerogel.

2.6. Transmission Electron Microscopy of Au~cyt. c Silica Aerogels

1. Au~cyt. c silica aerogel.

2.7. Nitric Oxide Sensing of Au~cyt. c Silica Aerogels

1. Au~cyt. c silica aerogel.

2. Cardboard platform (see Note 4).

2. PELCO® Center-Marked Grids, 200 mesh, 3.0  mm O.D., Copper (Ted Pella, INC). 3. Coat-Quick “G” grid coating pen (Ted Pella, INC).

2. Argon (or other inert gas), cylinder (Tech Air). 3. Small cylinder (i.e., 4 L) of 10% NO in argon (prepared inhouse) (see Note 5). 4. Tygon tubing (or other NO-resistant tubing), “T” switch valve, and syringe needles. 5. Disposable cuvette (Sarstedt, Acryl, 10 mm × 10 mm × 55 mm) with rubber septum cap.

3. Methods The cytochrome c heme protein remains viable throughout the conditions necessary to form an aerogel when in the presence of nucleating gold or silver nanoparticles. The model of the Au~cyt. c superstructure consists of the positively charged heme edge of the protein specifically adsorbing to the negatively charged citrate stabilized colloidal gold surface (13, 16–18). This highly curved surface presents the negatively charged side of the protein to the

Cytochrome c Stabilization and Immobilization in Aerogels

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medium in such a way that rapid (and likely weak) protein–protein association organizes the nonadsorbed protein into the multilayered superstructure (Fig. 1a) (13, 19). The organization of the superstructures is rapid and occurs in