STABILITY AND STABILIZATION OF BIOCATALYSTS
STABILITY AND STABILIZATION OF BIOCATALYSTS
Progress in Biotechnology Volume 1 New Approaches to Research on Cereal Carbohydrates (Hill and Munck, Editors) Volume 2 Biology of Anaerobic Bacteria (Dubourguier et al., Editors) Volume 3 Modifications and Applications of Industrial Polysaccharides (Yalpani, Editor) Volume 4 Interbiotech '87. Enzyme Technologies (Bla~ej and Zemek, Editors) Volume 5 In Vitro Immunization in Hybridoma Technology (Borrebaeck, Editor) Volume 6 Interbiotech '89. Mathematical Modelling in Biotechnology (Bla~ej and Ottov&, Editors) Volume 7 Xylans and Xylanases (Visser et al., Editors) Volume 8 Biocatalysis in Non-Conventional Media (Tramper et al., Editors) Volume 9 ECB6: Proceedings of the 6th European Congress on Biotechnology (Alberghina et al., Editors) Volume 10 Carbohydrate Bioengineering (Petersen et al., Editors) Volume 11 Immobilized Cells: Basics and Applications (Wijffels et al., Editors) Volume 12 Enzymes for Carbohydrate Engineering (Kwan-Hwa Park et al., Editors) Volume 13 High Pressure Bioscience and Biotechnology (Hayashi and Balny, Editors) Volume 14 Pectins and Pectinases (Visser and Voragen, Editors) Volume 15 Stability and Stabilization of Biocatalysts (Ballesteros et al., Editors)
Progress in Biotechnology 15
STABILITY AND STABILIZATION OF BIOCATALYSTS Proceedings of an International Symposium organized under auspices of the Worldng Party on Applied Biocatalysis of the European Federation of Biotechnology, the University of Cordoba, Spain, and the Spanish Society of Biotechnology Cordoba, Spain, April 19-22, 1998
Edited by A. B a l l e s t e r o s
Department of Biocatalysis, CSIC Institute of Catalysis, Madrid, Spain
F. J. P l o u
Department of Biocatalysis, CSIC Institute of Catalysis, Madrid, Spain
J. L. I b o r r a
Department of Biochemistry and Molecular Biology "B", Faculty of Chemistry, University of Murcia, Murcia, Spain P. J. H a i l i n g
Department of Pure & Applied Chemistry, University of Strathclyde, Glasgow, UK
(,
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Preface The aims of the Working Party on Applied bioeatalysis of the European Federation of Biotechnology are: i) to increase the understanding of bioeatalysis, in particular its commercial applications; ii) to take initiatives in areas of growing scientific and industrial interest and importance in the field of applied bioeatalysis; iii) to identify key topics which may be rate-limiting the development of scientific and technological capabilities in applied biocatalysis and to take steps to stimulate these areas. In the light of these aims, the Working Party has organised over a dozen symposia since 1985. In 1992 in Maastricht (The Netherlands) the WP held a symposium on Stability and Stabilization of Enzymes (ref. 1). We felt that six years later would be a good time to organize a second symposium. This would bring up to date the work already going on, and identify possible breakthroughs in the research. This time the scope of the conference was broadened by including whole cell biocatalysts in addition to enzymes. The international symposium on "Stability and Stabilization of Biocatalysts" was held in Cordoba, Spain from April 19-22, 1998. It was possible to bring together 210 participants, representing every continent, to learn from 150 oral and poster communications. Delegates had the opportunity to hear lectures on stability and stabilization grouped within the topics: inactivation mechanisms and reactors; stabilization by chemical modification; noncovalent processes in solution; protein engineering and thermophile enzymes; immobilized enzymes; non-conventional media; and whole cells. The papers included in these Proceedings are grouped within the above mentioned topics. We hope that the symposium, and this book, which contains most of the papers presented in Cordoba, will make a useful contribution to this key area of applied biocatalysis. We thank Professors J.M. Marinas and D. Luna, Department of Organic Chemistry, University of Cordoba, Spain, for their valuable contribution to the organization of the congress.
The Editors, Madrid, Murcia and Strathclyde, May 1998.
REFERENCE I. W.J.J. van der Tweel, A. Harder and R.M. Buitelaar (Eds.),Stabilityand Stabilization of Enzymes, Elsevier,Amsterdam, 1993.
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Acknowledgements The Organizing Committee gratefully acknowledges the support of the following sponsors: Diputaci6n de C6rdoba Ayuntamiento de C6rdoba Universidad de C6rdoba (Programa propio) Junta de Andalucia Comisirn Interministerial de Ciencia y Tecnologia (CICYT) Consejo Superior de Investigaciones Cientificas (CSIC) Instituto de Cat~ilisis y Petroleoquimica, CSIC Caja Murcia AVE-RENFE IBERIA Airlines Caja Sur
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Contents Preface Acknowledgements
vii
Inactivation mechanisms and Reactors Diagnosing the inactivating process of enzymes O. Misset, A. van Dijk Stability and stabilization of glucose-fructose oxidoreductase from Zymomonas mobilis against irreversible inactivation during substrate turnover in biochemical reactors B. Nidetzky, M. Ftirlinger, D. Haltrich, K.D. Kulbe
19
Reactor performance under thermal inactivation and temperature optimization with chitin-immobilized lactase A. Illanes, L. Wilson, C. Altamirano, A. Aillaphn
27
Fast in-situ-characterization ofbiocatalysts long-term stability M. Boy, A. Dominik, H. Voss
35
Chemical and thermal stability of ferulic acid (feruloyl) esterases from Aspergillus C.B. Faulds, F.O. Aliwan, R.P. de Vries, R.W. Pickersgill, J. Visser, G. Williamson
41
Stability properties of calf intestinal alkaline phosphatase U. McKeon, B. O'Connor, C. 0'Ffig~tin
47
Stabilization of [3-glucanase and its effect on the substrate hydrolysis pattern I. Markovi6, B. Markovi6-Dev~i6, S. Gamulin, N. Pavlovi6
53
Some factors affecting the behavior of anhydrous ~x-chymotrypsin at high temperature D. Pirozzi, G. Greco Jr.
59
Horseradish peroxidase stability in the course of phenols oxidation A.M. Egorov, Yu.L. Kapeluich, T.A. Pastuhova, M.Yu. Rubtsova, D.N. Cherepanov
65
The effect of impurities of crude olive residue oil on the operational stability of the Candida rugosa lipase immobilized in polyurethane foams A.C. Correia, S. Ferreira-Dias
71
Approaches for improved identification of mechanisms of enzyme inactivation M. Polakovi6, P. Vr~tbel, V. B~tle~
77
Stability and stabilization of a-l,4-D-glucan phosphorylases R. Grie[31er, B. Mtiller-Fembeck, S. D'Auria, F. La Cara, B. Nidetzky
83
Oxidation by hydrogen peroxide of D-amino acid oxidase from Rhodotorula gracilis V. Obreg6n, I. de la Mata, F. Ram6n, C. Acebal, M.P. Castill6n
89
Cosolvent effect on the synthesis of ampicillin and cephalexin with penicillin acylase C. Aguirre, J. Baeza, A. Illanes
95
Conversion and stability studies on enzyme-membrane reactor with lipase immobilized by different methods T. Venyige, E. Cshnyi, Cs. Sisak
101
Study of the deactivation process of the glucose oxidase-catalase enzymatic system by means of simulation L.E. Romero, D. Cantero
107
Stabilization by chemical modification
113
Stabilization ofhydrolases by chemical modification with fatty acids or polyethylene glycol F.J. Plou, M.V. Calvo, M. Ferrer, A. Ballesteros
115
Modification of the carbohydrate moiety in ribonuclease B and its influence on the protein stability probed by limited proteolysis U. Arnold, R. Ulbrich-Hofinann
121
Comparative studies on selective modification of e-amino groups in lipases and phospholipase A2 B.C. Koops, A.J. Slotboom, H.M. Verheij
127
Analysis of the thermal deactivation kinetics of a-chymotrypsin modified by chemoenzymatic glycosilation M.A. Longo, D. Combes
135
Increasing the operational stability of flavoproteins by covalent cofactor binding W.J.H. van Berkel, R.H.H. van den Heuvel, M.W. Fraaije, C. Laane
141
Properties of PEG-modified microbial proteases. Activity and stability studies T.M. Fatum, A. Agerlin Olsen, C.C. Fuglsang, D. Otzen
147
The effect of crosslinking on thermal inactivation of cellulases J. Bilen, U. Bakir
151
Studies of the stability of acid phosphatase (A. niger) by cross-linking with glutaraldehyde and soil humates N. Ortega, L. Berzal, M.D. Busto, M. Perez-Mateos
157
Non-covalent processes in solution
163
Folding and association versus misfolding and aggregation of proteins R. Jaenicke
165
Activity of monoclonal antibodies in prevention of in vitro aggregation of their antigens B. Solomon, T. Katzav-Gozanski, R. Koppel, E. Hanan-Aharon
183
Stability of monodeamidated forms of ribonuclease A F. Catanzano, G. Graziano, S. Capasso, G. Barone
189
Pressure effects on protein oligomeric dissociation C. Balny
197
Effect of high hydrostatic pressure on enzyme stability V. AthOs, D. Combes
205
Thermodynamic stability of a monomeric derivative of bovine seminal ribonuclease G. Barone, F. Catanzano, G. Graziano, V. Cafaro, G. D'Alessio, A. Di Donato
211
Thermodynamic stability of ribonuclease P2 sulfolobus solfataricus and some mutants G. Graziano, F. Catanzano, P. Fusi, P. Tortora, G. Barone
217
Stabilization of ct-chymotrypsin by DMSO E. Flaschel, L. Ebmeier
223
Thermal stabilization by its ligands ofNADP÷-isocitrate dehydrogenase from the thermophilic cyanobacterium Phormidium laminosum M.A. Pardo, M.J. Llama, J.L. Serra
229
The characterization of protein stability from hydrogen exchange: maximum entropy analysis of co-solvent effects on exchange rates B. Souhail, I.M. Plaza del Pino, J.M. Sanchez-Ruiz
235
Preferential hydration changes upon protein unfolding in water-cosolvent mixtures H.O. Hammou, I.M. Plaza del Pino, J.M. Sanchez-Ruiz
241
Charactbrizing protein-cosolvent interactions coupled to protein refolding by kinetic calorimetry M.C. Boulaich, A. Parody-Morreale, J.M. Sanchez-Ruiz
247
Characterizing cosolvent effects on protein stability: cold-denaturation of ubiquitin in the presence of guanidine B. Ibarra Molero, J.M. Sanchez-Ruiz
251
Acetylcholinesterase-ethanol interactions: inactivation, substrate and ligand exclusion F. Ortrga, D. Garcia, J.-L. Marty
257
xii Effect of a mixed stabilizer-denaturant system on the stability of enzyme activity at high temperatures: lysozyme as a model Y. Sangeeta Devi, U.B. Nair, R. Bhat
263
A simple folding method for high level production of the hydrophobic disulfide bonded hepatitis B X protein by inclusion body route and its structural analysis I. Marczinovits, J. Molnhr, M.Z. Kele, P.T. Szab6, T. Janfiky
269
Protein engineering and Thermophile enzymes
275
Rigidity of thermophilic enzymes A. Fontana, V. de Filippis, P. Polverino de Laureto, E. Scaramella, M. Zambonin
277
Improvement of thermal stability of a diagnostic enzyme, Streptomyces cholesterol oxidase, by random and site-directed mutageneses and a structural interpretation Y. Murooka, Y. Nishiya, M. Toyama, M. Aoike, M. Yamashita, N. Hirayama
295
Protein engineering for thermostabilisation of proteins: some theoretical rules and application to a 13-glucanase E. Querol, J. Pons, J. Cedano, M. Vallmitjana, F. Garcia, C. Bonet, J. P6rez-Pons, A. Planas, A. Mozo-Villarias j//
303
Mechanisms of stabilization of the [3-glycosidase from the hyperthermophilic archaeon
Sulfolobus solfataricus M. Moracci, M. Ciaramella, L.-H. Pearl, B. Cobucci Ponzano, M. Rossi
311
A strategy for engineering thermostability: the case of cyclodextrin glycosyltransferase J.C.M. Uitdehaag, B.W. Dijkstra
317
Thermophilic esterases and the amino acid "traffic rule" in the hormone sensitive lipase subfamily G. Manco, F. Febbraio, M. Rossi
325
Cloning and stabilization of NAD-dependent formate dehydrogenase from Candida boidinii by site-directed mutagenesis H. Slusarczyk, M. Pohl, M.-R. Kula
331
Usefulness of bacterial thermostable enzymes in clinical chemical analysis K. Tomita, K. Nomura, T. Miura, H. Shibata
337
Effect of temperature and Ca2÷ on the degree of multiple attack exhibited by mesophilic and thermophilic a-amylases A. Kramhott, B. Svensson
343
Degradation and denaturation of stable enzymes D. Thompson, R. Fernhndez-Lafuente, C. Mateo, D.A. Cowan, J.M. Guisfin, R. Daniel
349
xiii
Non-conventional media
353
Engineering stability of enzymes in systems with organic solvents V.V. Mozhaev
355
Inactivation of enzymes at the aqueous-organic interface P.J. Hailing, A.C. Ross, G. Bell
365
Exploiting hysteresis for high activity enzymes in organic media J.Partridge, P.J. Hailing, B.D. Moore
373
Limited proteolysis of proteins by thermolysin in trifluoroethanol P. Polverino de Laureto, E. Scaramella, M. Zambonin, V. De Filippis, A. Fontana
381
Enzyme inactivation by inert gas bubbling M. Caussette, A. Gaunand, H. Planche, B. Lindet
393
Effects of water-miscible solvents on the stability and specificity of cyclodextringlucosyltransferases A.D. B lackwood, C. Bucke
399
Stabilization of immobilized enzymes against organic solvents: complete hydrophilization of enzymes environments by solid-phase chemistry with polyfunctional macromolecules R. Femhndez-Lafuente, C.M. Rosell, L. Caanan-Haden, L. Rodes, J.M. Guishn
405
Effect of sorbitol on immobilized cx-chymotrypsin thermostability in low-water system T. de Diego, P. Lozano, M.J. lqiguez, J.L. Iborra
411
Stability of immobilized enzyme-polyelectrolyte complex against irreversible inactivation by organic solvents V. Levitzky, P. Lozano, A. Gladilin, J.L. Iborra
417
Enhancement of invertase activity in organic media for oligosaccharide synthesis S. Bielecki, R.I. Somiari
423
Effects of crown ethers on the activity of enzymes in peptide formation in organic media D.J. van Unen, J.F.J. Engbersen, D.N. Reinhoudt
429
Recovery of the activity of an immobilized lipase after its use in fat transesterification S. Ferreira-Dias, C.S. Duarte, V. Falaschi, S.R. Marques, J.H. Gusm~o, M.M.R. da Fonseca
435
Effects of lipid-bome compounds on the activity and stability of lipases in microaqueous systems for lipase-catalyzed interesterification X. Xu, C.-E. Hoy, J. Adler-Nissen
441
xiv Stabilisation of lipases for activity in ammoniolysis F. van Rantwijk, A.C. Kock-van Dalen, R.A. Sheldon
447
Water sorption isotherm as a tool to explore hydration of the microenvironment of biocatalysts J.M. Shnchez-Montero~ R.M. de la Casa, J.V. Sinisterra
453
Thermal stability of free and immobilised Pseudomonas cepacia lipase in aqueous and organic media G. Pencreac'h, J.C. Baratti
459
The stabilization of enzyme in organic solvent at low temperature H.J. Lee, J.R. Kim, Y.J. Yoo
465
A comparative study of thermal inactivation of enzymes in supercritical carbon dioxide A. Giel3auf, T. Gamse, E. Klingsbichel, H. Schwab, R. Marr
471
Studies of the stability of aminoacylase in some organic solvents L. Boross, J. Koshry, 1~. Stefanovits-Bhnyai, C. Sisak, B. Szajhni
477
Cutinase activity and enantioselectivity in supercritical fluids N. Fontes, M.C. Almeida, S. Garcia, C. Peres, J. Grave, M.R. Aires-Barros, C.M. Soares, J.M.S. Cabral, C.D. Maycock, S. Barreiros
483
Effect of pressure on enzyme activity in compressed gases M.C. Almeida, N. Fontes, E. Nogueiro, S. Garcia, C. Peres, A. Silva, M. Carvalho, S. Barreiros
487
Immobilized enzymes
493
Activity and structural stability of adsorbed enzymes W. Norde, T. Zoungrana
495
Covalent immobilization of glucose oxidase on A1PO4 as inorganic support. F.M. Bautista, M.C. Bravo, J.M. Campelo, A. Garcia, D. Luna, J.M. Marinas, A.A. Romero
505
The effect of site-specific immobilization on the thermal stability of thermolysin-like neutral proteases J. Mansfeld, G. Vriend, B. Van den Burg, G. Venema, V.G.H. Eijsink, R. UlbrichHofmann
513
Immobilization ofhydantoin cleaving enzymes from Arthrobacter aurescens DSM 3747 - Effect of the coupling method on the stability of the L-N-carbamoylase M. Pietzsch, H. Oberreuter, B. Petrovska, K. Ragnitz, C. Syldatk
517
Thermoinactivation of l~-xylosidase immobilized on nylon M.J. Duefias, P. Estrada
523
XV
Increase of thermal stability of tannin acyl hydrolase by covalent immobilization through its carbohydrate side chain: application in tea cream solubilisation P. Nicolas, J.L. Sauvageat, S. Reymond, E. Raetz
529
Stability in the presence of organic solvents of dextransucrase from Leuconostoc mesenteroides NRRL B-512F immobilized in calcium-alginate beads M. Alcalde, F.J. Plou, M.T. Martin, M. Remaud, P. Monsan and A. Ballesteros
535
Immobilisation and characteristics of glucose oxidase immobilised on Convective Interaction Media (CIM) disks A. Podgomik, M. Vodopivec, H. Podgomik, M. Barut, A. Strancar
541
Immobilization of spinach leaf hydroperoxide lyase L.M. Simon, Sz.J. Mfirczy, M. Kotorman, Sz.A. N6meth, B. Szajhni
547
A new immobilization system for Candida rugosa lipase: characterization and applications K. Carbone, M. Casarci
553
Soybean lipoxygenases: purification and stability of the free and immobilized enzymes A. Chikere, B. Galunsky, V. Kasche
559
Stabilization of lipase B from Candida antarctica by immobilization on different supports M. Arroyo, J.M. S~nchez-Montero, J.V. Sinisterra
565
Covalent immobilization of crude and partially-purified lipases onto inorganic supports: stability and hyperactivation A.R. Alchntara, I. Borreguero, M.T. L6pez-Belmonte, J.V. Sinisterra
571
Immobilization of alkaline phosphatases on various supports B. Surinenaite, V. Bendikiene, B. Juodka
577
Stabilization of serine proteases by immobilization V. Bendikiene, B. Juodka
583
Whole cells
589
Immobilized Cells: Plasmid stability and plasmid transfer J.N. Barbotin, D. Mater, M. Craynest, J.E. Nava Saucedo, N. Truffaut, D. Thomas
591
Stabilization of immobilized cells systems using a modified metal surface, fructose polymer levan and high cell concentration M. Bekers, E. Ventina, A. Karsakevich, I. Vina
603
Plasmid stability in recombinant Saccharomyces cerevisiae expressing the EXG1 gene in free immobilized cultures A. Guillhn, T. Lfi Chau, E. Roca, M.J. Nfifiez, J.M. Lema
611
xvi Bioreductions by Pyrococcusfuriosus at elevated temperatures E. van den Ban, H. Willemen, H. Wassink, H. Haaker, C. Laane
619
Stability of free and immobilized Mycobacterium sp. cells in aqueous and organic media P. Fernandes, J.M.S. Cabral, H.M. Pinheiro
625
Operational stability of immobilized C. reinhardtii cells: an approach to its potential as biocatalyst for N-consuming processes I. Garbayo, C. V ilchez
631
Induction and stability of cholesterol oxidase from cells of a Rhodococcus J. Kreit, P. Germain
639
Stability of functionally active fusion proteins during their biosynthesis and isolation from expressing bacterial cells L.M. Vinokurov, A.V. Yakhnin, T.V. Ivashkina, V.N. Ksenzenko, Yu.B. Alakhov
645
Miscellaneous Production ofbio-esters by immobilized lipases K. B61afi-Bak6, L. Gubicza, E. Cs~nyi, M.A. Galhn, J.A. Moreno
653
An aspergillusflavus strain promoting oleic acid esterification in isooctane V. Loscos, G. Caries, B. Perpifia, M. Torres, N. Sala, R. Canela
657
Covalent enzyme immobilization in different porous polymer membranes H.-G. Hicke, M. Becker, G. Malsch, M. Ulbricht
661
Functionalised cross-linked polyvinyl alcohol as new matrix for lipase immobilization E. Cernia, G. Milana, G. Ortaggi, C. Palocci, S. Soro
667
Enzymatic biphasic membrane reactors for the synthesis of chiral products H.A. Sousa, J.P.S.G. Crespo
673
Coimmobilization of enzymes and cells on chitosan and derivatives A.B. Martin, M. Picciolato, A. Heras
679
Parameters affecting the activity ofMucor miehei esterase 30 000 in a solvent freesystem M. Karra-Chaabouni, S. Pulvin, D. Thomas
685
Recombinant antigens by fusion of antigenic epitopes to a GST partner J. Molnhr, Marczinovits, M. Kiss, S. Husz, G. T6th, L. Dorgai, M. Khlmhn
691
Modification of metal substrates and its application to the study of redox proteins T. Pineda, J.M. Sevilla, A.J. Roman, R. Maduefio, M. Blazquez
697
xvii
Influence of some inducers on activity of ligninolytic enzymes from corncob cultures of Phanerochaete chrysosporium in semi-solid-state conditions S. Rodriguez Couto, M.A. Longo, C. Cameselle, A. Sanromhn
703
NMR study of hydration of liquid phase during lipase catalysed esterification in non aqueous media C. Sarazin, C. Roblot, B. Decagny, F. Ergan, J.N. Barbotin, J.P. Srguin
709
ot-chymotrypsin-catalysed synthesis of N-acetyl-L-tyrosine esters in organic media K. L~tszl6, L.M. Simon
713
Purification and characterization of penicillin V acylase from Streptomyces lavendulae R. Torres, I. de la Mata, M.P. Castill6n, M. Arroyo, J. Torres, C. Acebal
719
Studies on the regioselective acylation of sugars catalyzed by lipase in tert-butanol V. Sereti, H. Stamatis, F.N. Kolisis
725
Activity of cardosins A and B in the presence of organic solvents A.C. Sarmento, M. Barros, E. Pires
731
Effect of fermentation conditions in the enzyme activity and stereoselectivity of crude lipase from Candida rugosa R.M. de la Casa, A. SLnchez, J.V. Sinisterra, J.M. S~nchez-Montero
735
Biotransformations catalyzed by Candida rugosa lipase partially purified by precipitation and by organic solvents treatment S. Chamorro, A.R. Alchntara, J.M. S~.nchez-Montero, J.V. Sinisterra
741
Effect of pH and temperature on immobilised Thiobacillusferrooxidans cells over nickel alloy fibre J.M. G6mez, D. Cantero
747
Index of authors
753
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Inactivation mechanisms and Reactors
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editorg) 9 1998 Elsevier Science B.V. All rights reserved.
Diagalosing the inactivating process o f e n z y m e s Onno Misset and Alard van Dijk Research and Development, Food Specialties Division, Gist-brocades P.O. Box 1, NL 2600 MA, Delft, The Netherlands
1.
INTRODUCTION
Enzymes are widely used in a large variety of industrial applications (for an excellent text book see [1 ]). The commercial products appear in various formulations such as liquids, powders, granulates, tablets etceteras containing either single enzymes or mixtures of enzyme activities. An important aspect of enzymes is their stability, both during production and storage as well as their ultimate application. Elevated temperatures, extremes of pH, highand low salt concentrations and high water activity are known to be responsible for a low or reduced stability. Therefore, enzymes are produced and applied under conditions such as to retain maximal activity. Several methods are available to improve the stability properties of enzymes [2]. The use of stabilising additives is well-known and widespread and also protein engineering has proven to be successful in developing intrinsically more stable mutant enzymes. Chemical modification and immobilisation are, although also recognised as stabilising techniques, much less applied Despite the enormous amount of literature on stabilising additives, relatively little is known about the precise reasons why particular stabilisers work as good as they do. This is at least in part caused by the lack of knowledge about the actual enzyme inactivating processes. Also the success of protein engineering is significantly increased if the inactivating process is known at the molecular level. Of course some common inactivation mechanisms will play a role, but each enzyme has its own 'individual' sensitivities. Knowledge of these sensitivities can direct the stabilising process and contribute to the development of cost-effective enzyme products. In this article we will focus on how to identify the inactivating process. Hereto we propose a series of coherent experiments that is designed to quickly and unambiguously identify said inactivating event. This proposal emerged from extensive discussions between enzymologists, protein- and analytical chemists and formulation technologists from our company. We combined knowledge on enzyme stability (section 2) and the various analytical methods and their potential to be applied to enzyme products (section 3) in a multidisciplinary mode in order to arrive at proposal "diagnosing the inactivating processes of enzymes" (section 4).
2.
STABILITY OF ENZYMES
The stability of industrial enzymes is put to the test during all the various stages of its life as described earlier [2]. The first stage is the production process at the enzyme manufacturer's and comprises usually fermentation, downstream processing, formulation and storage. The subsequent stage involves transport and usage by the customer in a specific application. The enzyme molecules have to be able to survive all the different processing steps, each with its own physical and chemical conditions, in order to end up in the enzyme product, preferably at very high yields and then to deliver its promised performance in the customer's application. In order to do so, enzyme manufacturers are selecting and developing enzymes with already a high intrinsic stability and low sensitivity toward all kinds of inactivating processes. However, since enzymes still are protein molecules, they do possess a vulnerable three-dimensional structure with reactive amino acid side chains (acids, bases, nucleophiles etceteras).
~ non-covalent t
" 1 .., covalent
denaturation aggregation precipitation
oxidation deamidation Maillard proteolysis racemization hydrolysis
Figure 1. Scheme for the irreversible inactivation of an enzyme. N is the native enzyme; D represents an ensemble of reversibly, partially denatured forms.
The irreversible inactivation of an enzyme can be represented by the scheme in Figure 1 ([3] and reproduced from [2]). The enzyme molecule normally exists in its native, active state N which is in equilibrium with the partially denatured, enzymaticaUy inactive state D. At rising temperatures, the enzyme tends to unfold in a co-operative process. The temperature at which [N] = [D] is called the melting temperature Tin. The Tm of enzymes may vary from 30 to >100 ~ but, in aqueous solution is usually somewhere between 60 and 80 ~
Two different inactivating processes can be discerned: non-covalent and covalent. At higher temperatures, the unfolding process can proceed to such an extend that refolding is no longer possible because of denaturation accompanied by aggregation and precipitation. As a result, these non-covalent processes cause irreversible inactivation. The covalent processes involve chemical modification of the enzyme molecule because sensitive amino acids react with for instance oxidants, acids or bases, reducing sugars etceteras. Also included here is 'simple' proteolysis that may occur in case either the enzyme product is 'contaminated' with proteases (e.g. also produced by the production organism) or in case these are present in the application of the customer. The following physical and chemical factors are considered bad for enzyme stability because they either promote the non-covalent unfolding and/or aggregation/precipitation processes or enhance the rate of the covalent processes: 9 high temperature 9 extremes ofpH 9 extremes of ionic strength 9 high water activity
Solid- versus liquid enzyme preparations: solvent effects. The solution state of an enzyme has important effects on its thermodynamic stability: in solution the enzyme will have different stability characteristics than in the dry state. Drying is known to often cause structural changes, but fortunately most of them are reversible [4]. Enzymes are hygroscopic. In the dry form they absorb water. The stability of the enzyme is very much dependent on the moisture content and, at low contents, T m (or better Zg for solids) decreases with increasing content (Tg: glass transition temperature). In addition to an improved thermodynamic stability, the dry enzyme is also better resistant against microbiological degradation or chemical inactivation (except gas mediated inactivations like oxidation). From a stability point of view the dry state is therefore preferred for enzyme storage. Nevertheless, liquid enzyme formulations are still used because: 9 Cost aspect: evaporating water is very expensive! 9 The enzyme is inactivated by the drying process or, 9 The application requires a liquid formulation (costumer demand) The choice of solvent system of the liquid formulation is very important. Although water is the common solvent for enzymes, co-solvents are often added to improve the stability of the native protein. They are often added in high concentrations [5] and can either be other solvents (e.g. glycerol) or solids (e.g. sucrose) and all aim at reducing the water activity. Inactivation and changes in the protein molecule Irreversible inactivation of enzymes is always associated with a change in one or more of its basic properties as summarised in Table 1. In order to be able to diagnose the inactivating process, it is therefore essential to avail of analytical techniques that allow to monitor these changes preferably in a sensitive and unambiguous way. In the next paragraphs (section 2.2), the effect of the different covalent modifications will be discussed followed by a discussion of the various analytical techniques (section 3).
Table 1 - Basic properties of enzyme molecules. PROPERTY
DETERMINED BY
Structure primary secondary tertiary quaternary 'Molecular weight
Amino acid sequence Occurrence of alpha-helices and beta-strands/sheets 3-Dimensional folding Subunit composition Determined by the chemical composition of the protein molecule (i.e. amino acid composition plus cofactors, metal ions etceteras). .........
Charge overall
distribution
- Hydrophol~icity
2.1
Resultant of positive and negatively charged amino acid residues. Strongly determined by pH. Zero at pI (isoelectric point), overall negative when pH>pI and viceversa Location of the charged and/or polar side chains of amino acids on the surface of the protein molecule. Together with the hydrophobicity important for functional and physical properties of enzymes. Occurrence of amino acids with hydrophobic side chains at the surface of the enzyme molecule. Together with charge distribution important for functional and physical properties of enzymes. . . . . . . .
Covalent modifications
Many covalent changes in protein molecules have been described in the literature as being causative of inactivation: oxidation, deamidation, MaiUard reactions, racemization, proteolysis and polypeptide chain breakage (Figure 1). The likelihood that these processes occur depends on whether the reactive amino acid side chains are accessible to the reacting species or not. This means that in the enzyme in the N-conformation only surface-exposed side chains will react. In the partially denatured structures D, many more residues may become exposed and the irreversible inactivation may proceed at higher rates. Table 2 summadses the reactivity of the various amino acids and the effect of the covalent modifications on either the charge/polarity of the enzyme molecule or the molecular weight (M.W.) - see also Table 1. It is clear from Table 2, that most chemical modification cause changes in the polarity and charge of the enzyme molecule as well as its molecular weight. Yet, the extend and sign of the changes varies with the type of chemical modification.
Table 2. Reactivity of amino acids and effect On enzyme molecule A M I N O ACID
REACTIVITY
P R O P E R T Y OF ENZYME
none
none
like normal organic acids (formation of esters, reduction to alcohols potent nucleophile, acylation, alkylation, arylation, Maillard reactions
charge and M.W.
MOLECULE AFFECTED
aliphatic acidic
Ala, Val, Leu, Ile, Pro, Gly Glu, Asp
basic
Lys
charge and M.W.
charge and M.W.
His acylation, alkylation, electrophilic substitution oxidation Arg
polar
sulphurcontaining aromatic
Ser, Thr Asn, Gln Met, Cys Trp, Tyr, Phe
charge and M.W. can form heterocyclic condensation product with 1,2 and 1,3 dicarbonly compounds like alcohols deamidation alkylation oxidation electrophilic substitution oxidation, alkylation
polarity, M.W. charge polarity/charge and M.W. polarity/charge and M.W.
Effect on activity Covalent modification of an amino acid within the active site of the enzyme will certainly lead to an altered (lower) activity of the enzyme because the interactions between enzyme and substrate/product are affected. When the enzyme is not completely inactive, we can speak of a cripple enzyme. Whether this is the case depends on the position and the role of the amino acid in the enzyme molecule. Covalent modification of amino acid residues outside the active site usually has not an effect on the activity of the enzyme, but, the modifications may reduce the (thermo)-stability of the enzyme. This can be explained in terms of a shift in the equilibrium between the (modified) forms of N and D
2.1.1 Oxidation Oxidation of amino acids can occur by oxygen (02 - whether or not activated by metal ions), hydrogen peroxide or other stronger oxidants such as peroxy acids (peracetic acid). These reactions proceed via electrophilic addition or, more general, via radical addition. The latter makes the reaction rather pH-independent. The most sensitive amino acids are the sulphur containing cystein and methionine. Less sensitive are the aromatic amino acids tryptophane, tyrosine, histidine and phenylalanine. Table 3 shows that the oxidation products of these amino acids possess a different chemical structure which will disturb the enzyme structure and hence its (catalytic) properties.
Table 3. Oxidation of amino acids AMINO ACID OXIDATIONPRODUCT Cystein -CH2-SH
Methionine -CH2-CH2-S-CH3
TryPtophane
Cystine -CH2-S-S-CH2Cysteic acid ..... -CHz-SO~H. . . . Sulfoxide -CH:-CH2-SO-CH3 Sulfon -CH2-CHz-OSO-CH~ Oxindole-alanine N-formylkynurenine
OXIDANT
.
.
Oxygen H202 Performic acid . . . . . . H202 Chloramine T Performic acid
Histidine
H202 Iodine H202, Periodate, Iodine
Tyrosine
H~O2, Periodate,Iodine
The oxidised form of the enzyme can be regarded as a cripple form of the enzyme molecule. The change in molecular weight can be very small: the oxidation of one methionine toward its sulfoxide only increases its molecular weight by 18 dalton and leaves the charge almost unaffected. Oxidation of one cystein toward the cysteinic acid introduces one additional negative charge and increases the molecular weight by 54 dalton. 2.1.2 Deamidation
This reaction involves the hydrolysis of the uncharged amino acids glutamine (Gin) and asparigine (Asn) into the negatively charged glutarnicacid (Glu) and asparticacid (Asp). The molecular weight is affectedjust by the substitutionof a NH2-group by an oxygen-atom (l dalton). The reaction increases with increasing pH and temperature [6]. Since deamidation is a hydrolytic reaction, it may be expected to proceed much slower in powders although this will be dependent upon the hydration. Since the reactiondoes not require specialchemicals, it may occur during all stages of the life cycle of the enzyme. The effect of introducing the negative charge(s) in the enzyme molecule can be instability,inactivityor both.
2.1.3 Maillard reaction Browning of protein samples upon heating with sugars was first observed by Maillard in 1912 [7]. This reaction, also referred to as glycation of protein, is not only very important in the food industry, but also plays a role in ageing of proteins like the eye lens crystallins and in certain diseases such as diabetes mellitus. The chemistry of the Maillard reaction is very complex and not fully understood. Usually, the consecutive reactions are divided on the basis of product formation: early and advanced glycation products. The first step in the glycation process is the reaction between the aldehyde group of a reducing sugar (such as glucose) and the free amino group of a protein. The latter is either the alpha-amino group of the N-terminus of the protein or the gamma-amino group of the lysine side chain. The result is a still labile Schiff base which undergoes a rearrangement to form the more stable Amadori product. The Schiff base and the Amadori product are called the early glycation products. The steps following the ketoamine formation are not completely elucidated. A large number of reactions follow, resulting in complex carbohydrate-protein complexes. These structures can possess chromophoric (browning) and fluorophoric properties and are able to cross link protein chains [8]. These compounds are called advanced glycation end products. Also, chain breakage has been reported to occur as a result of the Maillard reaction. Other studies revealed that not only lysines undergo a reaction with sugars, but also tryptophanes and tyrosines can undergo modification with advanced products of the Maillard reaction (e.g. dialdehydes) [9] Conditions or compounds which favour the Maillard reactions are high(er) temperature, organic acids (such as sorbic acid). The reaction is supposed to involve the unprotonated aminogroup of lysine residues. This means that, with a pK~ of approximately 10, the reaction increases with increasing pH. As a result of the chemical modification of the enzyme molecule, both its molecular weight and charge distribution (and colour) will severely be affected, accompanied by both inactivation and a further decreased stability. 2.1.4 Racemization This process converts the (natural) L-amino acids to the D-form and can occur with each amino acid of an enzyme except glycines. It has been reported that only Asn and Asp are the most susceptible to racemization [10]. Furthermore, threonine and isoleucine contain an asymmetric carbon atom in their side chain which can, in principle, undergo racemization. The effect of racemization can be (partial) unfolding accompanied by loss of activity, although some authors consider that the contribution of racemization to thermoinactivation is negligible [11 ]. The process takes place preferentially at the extremes of pH and at high(er) temperatures.
2.1.5 Proteolysis Proteolysis is the process in which the enzyme molecule (i.e the substrate) is degraded by a protease. The sites of proteolytic attack are determined mainly by the specificity of the protease, but also involves flexible parts of the substrate which are located at the surface. Factors which promote proteolytic activity are the temperature and partially denaturing conditions for the substrate. The effect of other conditions such as the pH and the presence of metal ions are also depending on the type of protease. In general, the higher the concentration of both protease and its substrate, the faster the rate of proteolysis and, therefore, the rate of
10 inactivation. Proteolytic degradation usually leads to inactivation because the enzyme will be cut into small fragments.
2.1.6 Polypeptide chain breakage Apart from chain breakage by proteolysis, non-enzymatic breakage can occur at the extremes of pH extremes and elevated temperatures. Especially Asn-Gly bonds appear to be very sensitive. Effects similar to those after proteolysis.
3.
ANALYTICAL METHODS - WHAT ARE THEY WORTH?
There are various methods available which can be used to study the biochemical properties (see Table 4 for an overview). These can be categorised into a few distinct groups that will be shortly commented.
3.1 Amino acid analysis Two different methods are commonly applied in order to supply information about either the N-terminal amino acid sequence of the native enzyme molecule or (tryptic or CNBr-cleaved) fragments thereof or the quantitative amino acid composition. Comparison of the results with those for the non-inactivated enzyme may indicate if and where chemical modification occurred. Both methods are neither very accurate nor reproducible. Therefore, at the least they can only be indicative rather then very conclusive. For instance, quantitative amino acid analysis has a standard deviation of at least 10% which means that it is impossible to conclude for instance that 1 out of 5 methionines is chemically modified or similar reductions took place for other amino acids. 3.2 Electrophoresis Polyacrylamide gel electrophoresis (PAGE) is a very easy and therefore commonly performed experirnent. It can be carried out under several different conditions. In the presence of the surfactant sodium dodecyl sulphate (SDS-PAGE) the enzyme molecule becomes completely unfolded and coated with the negatively charged surfactant. Because of the constant charge/size ratio, the enzymes only migrate in the gel according to their (subunit) molecular weight. The changes in molecular weight that can be detected are in the order of >500 (compare with subunit molecular weight ranging from 15,000 - 50,000 for most industrial enzymes). Migration of enzymes in their native state (Native PAGE and Capillary Zone Eleetrophoresis) depends on both the charge and molecular weight. A change of even one charge (e.g. caused by deamidation, oxidation, Maillard reactions ...), gives already a significant change in nmning behaviour (as we observed also with protein engineering by which charged amino acid residues were mutated - not shown here). With isoelectric focusing (IEF) the enzymes migrate to the pH-value in the gel where their net charge is zero. Therefore, changes in charge earl also be detected very sensitively with IEF, provided that the enzymes remain soluble under these conditions and that the pI's are within the pH-range of the technique (2
Table 4 Inventory of protein chemical properties measured by various analytical methods M.W. Charge/Hydrophobicity
Structure
|
J i t
Chemical analysis
f
/
/
yes yes
yes
Amino Acid Sequencing Quantitative Amino Acid Analysi~
/
Electrophoresis Capillary Zone Electrophoresis Isoelectric focussing Native PAGE SDS-PAGE 2D-electrophoresis
yes
yes yes yes yes
yes yes yes yes yes
Chromatography Hydrophobie interaction Ion exchange Reversed phase Size exclusion
yes
yes
yes yes
yes yes
yes yes yes yes
yes
Spectroscopic techniques Circular dichroism Fluorescence spectrospcopy Fourier Transform Infrared Infrared/NFTIR Mass Spectroscopy Nuclear Magnetic Resonance Optical rotation dispersion Rarnan UV/VIS X-ray crystallography
Thermal analysis I
Differential Scanning Calorimetry
yes
yes
yes yes yes
yes
yes yes
yes
yes
yes
yes yes yes
yes
yes
yes
yes
12 2D-electrophoresis combines IEF and SDS-PAGE as first and second dimension respectively, thereby maximising the separating capacity.
3.3 Chromatography In addition to using it for the preparation of purified enzyme, column chromatography can also be applied in an analytical way to study chemical properties of protein molecules. Size exclusion chromatography (SEC; also named gel filtration) separates molecules on the basis of their size (and shape). For enzymes this means the Stokes' radius of the globular molecule. Although the solid phase of the column is made as inert as possible, too often aspecific interactions between enzymes and the matrix are encountered that result in an elution behaviour that deviates from the theory! This limits the applicability of this technique. Ion exchange- and hydrophobic interaction chromatography (IEX and HIC respectively) separate the enzyme molecules on the basis of the surface properties, i.e. the occurrence and distribution of charges and hydrophobic amino acids. A small sieving effect and some aspecific interactions are also present as a consequence of the structure of the resin. Reversed phase chromatography separates enzyme molecules in their denatured state usually in organic solvent systems. In this case the total chemical composition of the enzymes is determining the elution behaviour.
3.4 Spectrometric techniques Table 4 lists several spectrometric techniques that can be used to study one or more of the structural elements of enzyme molecules. The techniques require advanced, complex and expensive equipment and skilled people that are specialised in usually only one of these techniques. Also most techniques require pure enzyme in high amounts and at high concentrations. Nevertheless they have proven to be powerful tool in studying enzyme structural and also functional aspects and therefore remain in the biochemist' toolbox when stability problems have to be solved (see section 4). Without going to much in depth, the characteristics of most of the techniques are summarised in Table 5. Especially, the different variants of mass spectroscopy, in combination with some chromatographic procedure in advance (e.g. LC-MS) allow for an unambiguous identification of the type of inactivating mechanism that was active on which type of amino acid at which position. The reason for this is that the technique is able to determine molecular weights of intact and/or fragmented enzyme molecules with an accuracy of 1 Dalton!
3.5 Thermal analysis Of the various thermal analyses of enzymes, Differential Scanning Calorimetry (DSC) is most frequently used to determine the melting temperatures of enzymes T m in solution of solid state as well as the glass transition temperature Tg of solid formulations. This technique shows very elegantly the effect of stabilising additives or mutations or the Tm. However, also this technique requires for some experiments pure enzyme at high concentrations and high(er) amounts. Furthermore, also this technique requires advanced, complex and expensive equipment and skilled people that are specialised in interpreting the data measured.
13 Table 5 Comparison of spectrometric techniques to characterise enzyme structure and structural changes. TECHNIQUE
ADVANTAGE
DISADVANTAGE
UV/VIS spectroscopy
Easy to perform
Fluorescence
Sensitive to conformafional changes
Very global information Difficult to interpret Often not very sensitive to structural changes Only for soluble enzymes No information about underlying structural change Complex interpretation Only for enzymes < 40 kDa Only for soluble enzymes in aprotic buffers
NMR
FTIR
CD
(LC)-MS
High information density Very sensitive finger print of enzyme structure Very sensitive to structural changes . Semi-quantitative secondary structure information Su!table for solids and liquids Semi quantitative secondary structure information Very sensitive to structural changes Suited for both solid and liquid samples Information relatively easy to interpret Universal and selective detector of molecular species Low detection limit (very sensitive) Information easy to interpret Suited for both solid and liquid samples
Not very sensitive to small structural changes Limited number of buffers suited Information difficult to interpret Limited number of buffers suited
Removal Of effluent is necessary Limited number of buffers suited
14 4. R E S U L T S - S T R A T E G Y TO IDENTIFY THE P R O C E S S E S ENZYME INACTIVATION
THAT CAUSE
The strategy we propose resembles a decision tree and involves several consecutive stages (see Figure 2). Depending on the outcome of each stage, a subsequent stage will be entered in order to get the answer: what is the cause of enzyme instability in this particular case. ~tage !
yes
Obvious cause?
I ~176
Stage II
yes
Start of problem solution
Stage IV
v
Method: LC-MS
Method: 2D-electrophoresis
A minor change in MW? ConformaUonal change?
v
Determine cause of change Identify critical residues
A major change in MW? Change in charge state?
Stage III
J
yes v
Method: LC-MS & CD
Figure 2: Schematic presentation of stages that lead to the identification of the cause of enzyme instability.
Stage I - Is there a n o b v i o u s cause for the observed enzyme instability? The first step usually involves confirmation of an observed loss of activity and inspection of the sample. Then one has to evaluate whether there is an obvious cause for the loss of the activity. All measurements should be compared with the specifications of the product while using a "museum sample" of the enzyme product as the positive control. The following product properties have to be investigated: A. Activity of the product. How and by whom was the product instability identified? Can this be checked and verified?
B. Colour of the product. Did the colour of the product change? If it has become darker Maillard- or oxidation reactions may have taken place. C. Smell of the product. An off-smell may indicate growth of micro-organisms (infection = microbial instability).
15 D. Clarity of liquid products. Turbid solutions may indicate (protein) precipitation or growth of micro-organisms. E. pH of the product. In case the pH differs from the original value, degradation of the enzyme by e.g. deamidation or proteolysis may have occurred.
F. Conductivity of the liquid product The answers to question A-F may either give a lead for direct problem solution (the so-called obvious case) for instance by better controlling the manufacturing, transport and/or storage of the enzyme product. But in those cases where all these properties of the enzyme product mentioned under B-F are unaffected (i.e. with the exception of the enzyme activity = A), we are entering into stage II in which we will focus on the enzyme molecule itself.
Stage H - Is there a major change in molecular weight and~or charge state of the protein ? In section 2, several processes leading to loss of activity were discussed. These were divided into two main categories: 9 Covalent modifications 9 Non-covalent modifications Covalent modifications invariably lead to a change in either the molecular weight of the enzyme molecule, its charged state or a combination of both; stage II is meant to identify these changes. The most straightforward way to do this is by 2-D gel electrophoresis.. This experiment will directly indicate any (major) change in either molecular weight (within resolution of the gel, i.e. >500 Dalton) or isoelectric point (theoretical calculations show that one charge mutation changes the pI of an enzymes by 0.10-0.15). A complication with industrial enzymes is that the enzyme of interest may be a minor component in the enzyme mixture. This means that a 2D-gel may contain several spots and it can be difficult if not impossible to identify the interesting spots. In some cases this problem can be solved by using enzymograms for detection of specific activities (e.g. gelatin test for proteases). However, inactivated species or enzyme fragments will not show up in such staining methods, making the method of limited use. The solution is to have polyclonal antibodies against the protein of interest that make it possible to identify the relevant protein (or protein fragments). Such an enzym-specific 'staining' is crucial in the problem analysis and requires: 9 Purified enzyme of interest 9 Preparation ofpolyclonal antibodies against the purified enzyme of interest 9 A reference 2D-pattern of the intact (purified) enzyme of interest The development of these relatively simple tools is essential for a quick and thorough analysis of the causes of enzyme instability. Purification of the enzyme of interest is essential in the other stages as well.
Conclusion r r
In stage II the enzyme product will be analysed by 2D-gel electrophoresis (IEF and SDS-PAGE) using regular or specific (enzymogram or antibody) staining. Antibody staining requires purification of the enzyme and antibody production
16 In case stage II indicates no covalent modifications of the enzyme of interest (within the resolution of the techniques described), the analysis enters stage III where smaller covalent changes or changes in the conformational properties of the enzyme can be detected. In the opposite case where a clear covalent modification can be demonstrated, the analysis enters stage IV in order to further unravel the precise nature of the chemical modification. Stage III- Is there a minor change in molecular weight or a conformational change in the protein with no mass changes? A small change in molecular weight arising from limited proteolysis (a few amino acids N- or C-terminal) or an electrically 'neutral' chemical modification (oxidation of methionine to the sulfoxide form) still can have occurred but undetectable by the method applied in stage II. As a result of its high sensitivity and high resolution (dalton range) mass spectroscopy is the method of choice to look at this range of minor modification. Alternatively, the enzyme instability is caused by a loss of the 3-dimensional conformational integrity of the enzyme molecule. This can be detected by methods such as nuclear magnetic resonance (NMR), Fourier transform infrared spectroscopy (FT-IR) and preferred by us - circular dichroism (CD). These analytical tools can only be applied to purified enzymes samples, i.e. single components in a defined buffer system. This requires: 9 Purification of the enzyme of interest 9 A reference spectrum the (unmodified) enzyme of interest In some cases a useful reference spectrum may be recorded of the crude sample but in the majority of cases the matrix will interfere.
Conclusion r r r r
Analysis in stage III requires purification of the enzyme of interest from both the inactivated as well as the reference product followed by, Recording of a mass spectrum to identify small changes in molecular weight and to identify critical residues (of peptides) Recording CD (optionally NMR or FTIR) spectrum and comparison with reference spectrum of unmodified reference enzyme Determination of Tin
Stage IV. Determination of the cause of the activity change and the identification of critical residues involved. It has been concluded in stage II that there is a covalent modification leading to enzyme instability and/or inactivation. In stage IV we will now focus on the question: what type of covalent modification (proteolysis, deamidation, oxidation, (de)glycosylation, Maillard) and optionally - what is (are) the sensitive site(s) in the amino acid sequence. The most straightforward analysis is mass spectrometry. This requires at least a partially purified enzyme often in combination with a peptide map of the enzyme [12].
Proteolysis Multiple bands on SDS-PAGE indicate proteolysis; purification of the fragments followed by mass spectroscopy, directly gives information about the sensitive sites in the protein, provided that the amino acid sequence is known (for the majority of industrial enzymes this is the case). Proteolysis can be caused by a contaminating protease or, in case
17 the enzyme of interest is a protease: autodigestion. These two options can be distinguished by diluting the sample and follow the rate of inactivation as a function of the dilution factor. In case of autodigestion the rate will be independent of enzyme concentration (first order process), in the other case it will be concentration dependent (second order process).
Modifications, other than proteolysis, leaving a soluble enzyme. In case of other types of covalent modifications, assuming the modified protein is still soluble, the modified protein will probably still behave similarly to the intact protein in chromatographic procedures, and can be purified accordingly. Covalent modifications all give rise to specific changes in MW; therefor, mass analysis of the purified protein (with or without peptide mapping) will directly give information about the type of modification. Deamidation, for instance, gives rise to a change in MW of only 1 dalton whereas oxidation and Maillard reactions will give rise to larger changes in MW, the size of which characterises the specific event (the MW change for the oxidation if two thiols to a disulphide differs from that for the oxidation of a thiol to a sulfoxide). Identification of the site of modification requires knowledge of the amino acid sequence of the protein; it can be determined using MS-MS analysis (two coupled mass spectrometers, separated by a collision chamber) of the intact protein or of MS analysis of the peptide fragments after enzymatic or chemical cleavage and purification. The inactivation process may be an inherent protein property or be catalysed by a matrix component; the two options can be distinguished via a dilution series, as described above. Modifications, other than proteolysis, resulting in insoluble protein. In case the modified protein becomes insoluble (e.g. due to aggregation), it will first have to be solubilised using chaotropic agents (urea, guanidine chloride) or detergents after which the same track can be followed as described above for the soluble modified protein. The purification procedure, however, will probably differ from the one for the native protein. Conclusion r Analysis in stage IV requires purification of the enzyme of interest from both the inactivated as well as the reference product followed by: r Mass spectroscopy: conclusion about the identity of the major covalent changes r Fragmentation of enzyme (peptide mapping) r Determination of sensitive site after fragmentation r Determination whether the inactivation process is an inherent enzyme property
5.
ACKNOWLEDGEMENT
We like to acknowledge our colleagues Ruud Barendse, Koos Batist, WiUem Bijleveld, Raoul Cassaigne, Wim Dusoswa, Jan-Metske van der Laan, Gabrie Meesters, Erry Oosterom, Marcel Pi~t, Dick Schipper, Mirjam Scholte, Paul Schuurhuizen, Marcel van Tilborg and Bert Vroemen from Gist-brocades for providing experimental data, reviewing the literature, stimulating discussions and financial support.
18 References
[ 1] Godfrey, T and West, S.I. (eds), Industrial Enzymology, Stockton Press, New York, 1996 [2] Misset, O. in Stability and stabilisation of enzymes, Studies in Organic Chemistry, 47, (1993) 111-131 (Tweel, W.J.J. van den, Harder, A. and Buitelaar, R.M. eds), Elsevier Science Publishers, Amsterdam [3] Lumry, R. and Eyring, H., J. Phys. Chem., 58, (1954), 110. [4] Griebenow, K. and Klibanov, A.M., Proc. Nat. Acad. Sc., 92, (1995), 10969-10976 [5] Timasheff, S.N. in Methods in Molecular Biology, 40, 253-269 (Shirley, B.A. eds), Humana Press Inc., Totowa (1995) [6] Aher, T.J. and Klibanov, A.M., Science, 228 (1985), 1280 [7] Maillard, L.C., C.R. Seances Acad. Sci. 154, (1912), 66-68 [8] Yen, G.C. et al. Food Sci. Technol. 29, (1989), 273-288 [9] Ashoor, S.H. et al. J. Food. Sci. 49, (1984), 1206-1207 [ 10] Zhao, M. et al. Bioorganic Chemistry, 17, (1989), 36-40 [ 11 ] Zale, S.E. and Klibanov, A.M. Biochemistry, 27 (1986), 5432-5444 [12] Williams, K.R. and Stone, K.L., Mol. Biotechnol. 8, (1997), 155-167
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
19
Stability and stabilization of glucose-fructose oxidoreductase f r o m Zymomonas
mobilis a g a i n s t i r r e v e r s i b l e
inactivation
during
substrate
turnover
in
b i o c h e m i c a l reactors B. Nidetzl~, M. F'tirlinger, D. Haltrich and K. D. Kulbe Division of Biochemical Engineering, Institute of Food Technology, Universitiit fur Bodenkultur Wien (BOKU), Muthgasse 18, A- 1190 Wien, Austria 1. I N T R O D U C T I O N Gluconic acid and sorbitol are chemical commodities with many applications in food and chemical industries. Both compounds are industrially produced from glucose whereby gluconic acid is derived by chemical or microbial oxidation and sorbitol is obtained by chemical reduction. The ethanologenic bacterium Zymomonas mobilis produces an enzyme, glucosefructose oxidoreductase (GFOR), that can synthesize both products in a simultaneous manner, thus converting a substrate solution equimolar in glucose and fructose into products [ 1,2]. The simultaneous utilization of glucose and fructose for production of, respectively, gluconic acid and sorbitol is thought to add significant value to the educt mixture, even though separation of the products is required after exhaustive substrate conversion. GFOR contains tightly bound cofactor, E-NADP(H), and catalyses two individual half reactions [3]: (i)
D-fructose + E-NADPH <-> D-sorbitol + E-NADP
(ii)
D-glucose + E-NADP <-> D-glucono-1,5-1actone + E-NADPH
(ii)'
D-glucono-1,5-1actone + H20 <-> D-gluconic acid
D-Glucono-1,5-1actone is unstable in aqueous solution at around neutral pH and hydrolyses spontaneously. The net reaction is a transfer of reduction equivalents from glucose to fructose mediated by enzyme-bound NADP(H), and the cofactor cycles between its oxidized and reduced form. GFOR seems to be specific for the acceptor molecule, fructose, but not for the donor aldose. As consequence, several aldose sugars other than glucose such as xylose, galactose and even lactose [4] can be oxidized into the corresponding aldonolactone, subsequently hydrolysing into aldonic acid. Compared with conventional coenzyme-dependent enzyme systems, one major advantage of GFOR is the activity of the enzyme without exogenous NADP(H), which is an expensive and quite unstable molecule. Relative to free NADP(H) that displays a half-life of no more than about 24 to 72 h [5], the GFOR-bound
20 NADP(H) seems to be significantly stabilized to chemical decomposition. Furthermore, binding to GFOR increases the apparent molecular mass of NADP(H) to about 41 kDa, compared with 0.7 kDa for free NADP(H). Use of macromolecular coenzyme greatly assists continuous process development. Retention of small molecules such as NADPffl) in immobilized enzyme reactors or ultrafiltration membrane reactors is not easily accomplished [6], and coenzyme losses will add to coenzyme-specific product costs. Especially for low-price high-volume products such as gluconic acid the contribution of coenzyme-specific product costs to total product costs could be critical. For conversion of glucose and fructose two different forms of GFOR could be useful biocatalysts: permeabilized cells of Z. mobilis and cell-free (partially) purified GFOR. Permeabilization of the bacterial cells was shown to yield a stable and apparently fully active biocatalyst [7]. In comparison with permeabilized Zymomonas the advantage of cell-free GFOR could be the utilization of higher enzyme activities per unit reactor volume and thus a significant increase in productivity. However, compared with stability of GFOR encapsulated by the cellular matrix of Z. mobilis, the operational stability of the isolated enzyme was shown to be low, even when a reaction temperature of only 25 ~ was employed [8]. The inactivation reduced the total turnover number for GFOR drastically and hampered efficient use of the isolated enzyme in biochemical reactors [8]. In contrast, a continuous reactor loaded with permeabilized cells was operated at 39~ for several days without apparent decrease in enzyme activity [7]. Interestingly, inactivation of free GFOR seemed to be coupled to substrate turnover because the enzyme was stable under otherwise identical incubation conditions when no catalytic action occurred [8]. Autoinactivation during catalysis (as with glucose oxidase 191) or product-induced inactivation of GFOR are interpretations that seemed consistent with the experimental observations. Our recent studies aimed at elucidating the mechanism of GFOR inactivation in some detail [ 10] and, based on this mechanistic understanding, improving the operational stability of the enzyme during substrate conversion [11]. Some of the results are reviewed here.
2. INACTIVATION OF GFOR DURING SUBSTRATE TURNOVER Judging from the full stability of GFOR at 25 ~ in the absence of turnover for at least 24 h, rapid inactivation during substrate conversion was a surprising result (Figure 1, A). The final extent to which enzyme activity was lost during the reaction was dependent on initial substrate concentration. A correlation between catalytic activity and inactivation was proposed [8] because remaining GFOR activity was stable when conversion of substrate had been completed in batch reaction. At high concentration of glucose and fructose, however, the fast depletion of enzyme activity with time did not allow complete substrate conversion (Figure 1,
21 A), unless fresh enzyme was added to restart the reaction. The inactivation of GFOR was irreversible and did not depend on one particular degree of purification of the enzyme. Hence, a negative effect of contaminating proteins or enzyme activities seemed unlikely.
~-
lOO
"~ ",1=:1
8o
c~
60
r~
40
N
20
DTr~
0
~
o 0
20
40
60
80
100
Incubation time (h) Figure 1. Stability of GFOR during substratc turnover at 25 ~ using 1 M glucose and fructose as substrates (according to ref. [8]). (A) Enzyme activity without addition of DTr. (B) Enzyme activity when 10 mM DTT was added during the reaction. 2 M KOH was used for pH control (pH 6.5 in 0.1 M potassium phosphate buffer). We tried to derive a simple mathematical model that would give a satisfactory description of enzyme inactivation as function of substrate conversion. An apparent kinetic correlation between rate of reaction and rate of enzyme inactivation was found, that is d[S]/dt = k- d[E]/dt, where [S] and [El are substrate and enzyme concentration, t is the reaction time and k is an empirical parameter that lumps the effects that lead to inactivation of GFOR [8]. Although the model is capable of describing well the time-dependent product formation including enzyme inactivation [8,11], the resulting values for k seem consistent only for a limited range of experimental conditions (Table 1). One major drawback of the model is that it cannot account for the marked substrate concentration-dependence of the rate of GFOR inactivation [8], and
further analyses have shown thatindce~ high concentrationsof fi'uctoseratherthan glucose are
responsiblefor stabilizationof G F O R activity(Table I).
Itwas found empiricallythattwo factorsother than high fructoseconcentration crucially
determine the operational stabilityof G F O R during substrateconversion in batch or continuous
reactors [I 1]: thiolprotectionwith dithiothreitol(DTr) in concentrationof 5 to 10 m M being the most efficientreagent and use of weak bases such as carbonate, "Irisor Imidazole rather than hydroxide for p H control, i.e.,neutralizationof gluconic acid (Table 1). Note that D T r was
22 effective only when present during the conversion. Supplementation of DTT to partially inactivated GFOR failed to restore enzyme activity anew, but slowed the loss of remaining activity during further substrate turnover (Figure 1, B). The operational stability of GFOR was good and thus efficient use of enzyme activity for product synthesis could be made when GFOR was retained by a 10-kDa ultrafiltration membrane in a continuously operated tank reactor with constant discharge and 10 mM DTI" and 2 M Tris (as base) were employed (of. Table 1). Under these conditions a stable conversion could be maintained for about 500 h. The corresponding loss of enzyme activity was no more than about 20%. Table 1. Rate-dependent inactivation of GFOR during substrate turnover in a continuous ultrafiltration membrane reactor under different conditions at 25 ~ (according to ref. [11]). i
(2 M base)
Thiol stabilisation Inactivation with DTI' constant (k x 102)
Half-life of GFOR activity (h)
0.1 / 0.1
KOH
10mM
0.180
= 100
0.5 / 0.1
KOH
10 mM
0.100
> 200
1.0 / 1.0
"Iris
10 mM
0.003
> 500
1.0 / 1.0
"Iris
none
0.012
- 200
Substrate
pH control
(M Fru / M Glc)
i
3. M U L T I S T E P MECHANISM OF GFOR INACTIVATION 3.1. Destabilizing effect of the lactone product
The stability of GFOR was determined in the presence of fructose, glucose, sorbitol or sodium gluconate (0.5 M each). Compared with the control, i.e., GFOR in buffer, none of these compounds seemed to negatively affect long-term stability of the enzyme at 25 ~
As
consequence of this result, both the NADPH- and the NADP-form of GFOR are considered to be stable enzyme forms. The effect of glucono-1,5-1actone on GFOR stability was difficultly quantitated because of the rapid hydrolytic decomposition of this compound into gluconic acid in aqueous solution at about neutral pH. Using the alternate product xylono-1,4-1actone (100 mM) that is less prone to hydrolysis than glucono-1,5-1actone (note that xylose is substrate for the oxidizing half reaction of GFOR), time-dependent irreversible losses of enzyme activity were observed [10]. Since pH changes (due to formation of xylonic acid) were very small in this reaction, destabilization of GFOR seems to be caused only by interactions of the enzyme with the lactone product. It is important to note, however, that excess of exogenously added
23 gluconolactonase, i.e., the enzyme that catalyzes the hydrolysis of glucono-1,5-1actone, could not stabilize GFOR activity during the conversion, hence in the presence of lactone. 3.2. Time-course of inactivation The time-dependence of GFOR inactivation provides some insight to the underlying mechanism. It was found that the loss of GFOR activity with reaction time was preceded by a characteristic lag time (Figure 2). The lag time was dependent on fructose concentration and increased with increasing fructose. This result suggested a series mechanism of inactivation in which the native enzyme, N, reacts to give one or several intermediate enzyme forms, [N*], with destabilized conformation relative to N, but fully retained enzyme activity. [N*] inactivate(s) irreversibly to give I, thus a plausible model is: N <-> [N*] -> I
(1)
According to model (1), binding of the lactone product to GFOR would trigger the N -> [N*] transition. It is interesting to note that in addition to high fructose concentrations thiol protection using 1 mM DTI" could significantly increase the lag time prior to the onset of inactivation (Figure 2).
~ v,,,~
100
o
80 6o
~
40
"~ 20 ~ ~,,,4
o
0 0
10 Reaction time (11)
20
Figure 2. Inactivation of GFOR during turnover is preceded by a lag period (according to reL [ 10]). Pure GFOR (0.5 I.tM) was incubated at 30~ in 50 mM phosphate-buffer, pH 6.4, in the absence or presence of substrate: El, no substrate; II 1.5 M substrate, 9 0.5 M substrate, O 0.5 M substrate and 1 mM Dq~F. Control of pH was with 2 M Tris, and the substrate concentration was kept constant using pH-controlled feed of glucose and fructose. 3.3. Involvement of reactive thiols
The effect of D'I~ for stabilizing enzyme activity suggests that reactive thiols may be involved in the inactivation. To ascertain a putative role of thiols for stability of GFOR, chemical modification of the enzyme was carded out using N-ethyl-maleimide fNEM) which is
24 a reagent quite specific for protein sulfhydryl groups. The modification reaction was carded out in the absence and presence of substrates. In each case the time-dependence of enzyme inactivation in the presence of NEM obeyed an apparent fast-order reaction for which a plot of the natural logarithm of residual enzyme activity vs. incubation time is linear. As shown in Figure 3 that compares the inactivation constants for different reaction conditions, GFOR was most rapidly inactivated by NEM when glucose and fructose were present at the same time, hence when catalysis occurred and lactone product was formed. Compared with the results obtained for GFOR in buffer, glucose had a weak destabilizing effect whereas fructose could even afford slight protection to the enzyme. The pH change because of gluconic acid formation was negligible for the reaction time studied, and GFOR was stable in the absence of NEM.
m
~
.o 6
..-~4 O
~,~
2 0 Buffer
Glc
Fruc
Glc/Fruc
Figure 3. Inactivation of GFOR as result of chemical modification of thiols using NEM. GFOR (0.3 gM) was incubated with 20 mM NEM in 50 mM potassium phosphate buffer, pH 7.3, at 25~
in the absence or presence of substrates (1 M each). The inactivation constant is the
negative slope of the plot In (residual enzyme activity) vs. incubation time. The results in Figure 3 implicate exposure and deactivation of reactive thiols in the inactivation mechanism of GFOR, especially during substrate turnover. The exposure of sulfhydryl groups seems to be the result of conformational changes of GFOR, thus probably reflecting the N -> [N*] transition induced by the lactone [10]. Because the reactive form of the sulfhydryl group is the thiolate anion that is favored at high pH, use of strong bases for pH control in line with local pH inhomogeneities due to imperfect mixing of the reactor volume is expected to increase the sensitivity of GFOR to thiol deactivation with subsequent loss of enzyme activity, which was observed (Table 1 and ref. [ 11]).
25
3.4. Aggregation determines the loss of enzyme activity A line of experimental evidence suggested that thiol deactivation is not directly responsible for loss of enzyme activity. First, GFOR previously inactivated by chemical modification using methylmethane-thiosulfonate (MMTS) which is a reagent that forms mixed disulfides with protein thiols in a readily reversed manner could not be re-activated by treatment with excess of DTT. Second, colorimetric titration of reactive thiols in N and [N*] with Ellman's reagent did not lead to immediate loss of enzyme activity [10]. Third, using light scattering as probe of the extent of protein aggregation in samples taken from a typical batch reaction, evidence for a time-dependent increase in formation of high-order protein associates was found. Protein aggregation was irreversible and was correlated kinetically with enzyme inactivation [ 10]. Model (1) should therefore be extended to, N <-> [N*] -> [N*]deact-> I
(2)
where [N*]deact is an intermediate with deactivated thiol residues but retained enzyme activity. [N*]deact is expected to be prone to formation of high-order protein associates and will thus aggregate irreversibly. Recent analysis has shown that thiol inactivation in GFOR proceeds sequentially and, by measuring surface hydrophobicity as function of reaction time, provided an explanation for the high tendency to aggregation of intermediate [N*]deaet [10]. 4. S T A B I L I Z A T I O N OF G F O R Improved performance of isolated GFOR during conversion in a continuously operated ultraf'fltration membrane reactor has been reported recently, and stabilization of enzyme activity was based on the empirical finding that use of DTF together with a weak base for pH control can confer acceptable operational stability to the free enzyme [11 ]. Results of mechanistic studies provide now a clearer explanation to these observations and allow to devise other and probably more efficient strategies to maintain GFOR in an active, non aggregated state I I01. Because fructose and lactone product compete for binding to the NADP-form of GFOR, it seems reasonable to assume that high concentrations of fructose displace gluconolactone from the binary GFOR-gluconolactone complex and thus stabilize N relative to N* during turnover. The difference in the equilibrium dissociation constants [12] of GFOR.fructose (> 1 M) and GFOR.gluconolactone (0.35 mM) indicates that fructose can only poorly compete with gluconolactone for binding to GFOR, and thus a very large excess of fructose would have to be present at all times. Thiol protection on the other hand will stabilize [N*] rather than N. Because the deactivation of exposed thiols did not immediately lead to inactivation, prevention of aggregation seems to be most important in order to avoid GFOR inactivation. Use of bovine serum albumine (BSA; 5 g/L) as anti-aggregation reagent could indeed stabilize well GFOR
26 activity during conversion [6]. The effect of BSA on reducing the aggregation of GFOR could unfortunately not be determined by light scattering because the presence of a large excess of other protein interfered with the measurements. Finally, we discuss whether the results described here can explain why GFOR is stable when encapsulated by the cell matrix of Z. mobilis [7]. The destabilizing interaction of the enzyme with glucono- 1,5-1actone can probably
not be avoided in the eeUular reaction system as such, even though a periplasmic gluconolactonase is known to be present in Z. mobilis. The entrappment of GFOR could however reduce the conformational flexibility of the enzyme in such a way that structural transitions observed with free GFOR are efficiently prevented (of. model (2)). The results with soluble enzyme suggest that avoidance of aggregation of [N*]deaet would be sufficient to prevent the enzyme inactivation.
REFERENCES o
U. H. Chun and P. L. Rogers, Appl. Microbiol. Biotechnol., 29 (1988) 19.
2.
M. Zachariou and R. K. Scopes, J. Bacteriol., 167 (1986) 863.
3.
M. Hardman and R. K. Scopes, Eur. J. Biochem., 173 (1988) 203.
4.
M. Satory, M. Fiidinger, D. Haltrich, K. D. Kulbe, F. Pittner and B. Nidetzky,
o
Biotechnol. Lett., 19 (1997) 1205. Chenault and G. M. Whitesides, Appl. Biochem. Biotechnol. 14 (1987) 147.
6.
K. D. Kulbe and H. Chmiel, Ann. N. Y. Acad. Sci., 542 (1988) A.'!.'!..
7.
B. Rehr, C. Wilhelm and H. Sahm, Appl. Microbiol. Biotechnol. 35 (1991.) 144.
8.
D. Gollhofer, B. Nidetzky, M. Ftirlinger and K. D. Kulbe, Enzyme Microb. Technol.
o
17 (1995) 235. C. Bourdillon, C. Hervagault and D. Thomas, Biotechnol. Bioeng., 27 (1985) 1619.
10.
M. Fiirlinger, D. Haltrich, K. D. Kulbe and B. Nidetzky, Eur. J. Biochem., 251
11.
(1998) 955. B. Nidetzky, M. Ftirlinger, D. Gollhofer, D. Haltrich and K. D. Kulbe, Biotechnol. Bioeng. 53 (1997) 623.
12.
M. Hardman, M. Tsao and R. K. Scopes, Eur. J. Biochem., 205 (1992) 715.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
27
Reactor p e r f o r m a n c e under thermal inactivation and temperature optimization with chitin-immobilized lactase A. IUanes, L. Wilson, C. Altamirano and A. Aillapfin Escuela de Ingenieria Bioquimica, Universidad Cat61ica de Valparaiso. PO Box 4059, Valparaiso, Chile
ABSTRACT Kinetics of thermal inactivation of chitin-immobilized lactase (CIL) was studied in the absence and presence of catalytic modulators. Results were in good agreement with two-phase series mechanism of inactivation. The inhibitory product galactose was a positive modulator of enzyme thermal stability, while the opposite held for the substrate lactose. Differential equations were developed and solved to describe packed-bed reactor operation with CIL, considering lactose and galactose modulation. The model was experimentally validated in a laboratory packed-bed reactor. A more stable CIL has been developed and their kinetic (Vm~, Kin, KI) and inactivation parameters (kd have been determined as explicit functions of temperature. A strategy for temperature optimization of CIL reactor operation is presented in which the experimental information gathered has been included.
1. INTRODUCTION Enzyme thermal inactivation, although a well reported phenomenon, has seldom been studied in the presence of catalytic modulators to develop sound models to describe enzyme reactor performance [1-2]. Thermal inactivation in the presence of catalytic modulators has been described already and characterized in terms of modulation factors, defined in equation 2 [3]. Substances that interact with the enzyme during catalysis are potential modulators of its thermal stability and, therefore, modulation factors are relevant parameters for evaluation of reactor performance. Chitin immobilized lactase (CIL) was used as a model enzyme; in this case, both the substrate lactose and the inhibitory product galactose are potential modulators. In fact, galactose was proven to be a positive modulator of CIL stability while lactose had the opposite effect [4]. Kinetic parameters (kcat, Km and KI) and inactivation parameters (kij) are all temperaturedependent. Therefore, temperature explicit functions for all of them will allow to develop a model for reactor temperature optimization.
28 2. THEORETICAL BACKGROUND
2.1. Enzyme inactivation Enzyme thermal inactivation can be conveniently modeled based on a series type mechanism [5]. Reaction kinetics and enzyme inactivation for CIL according to a two-stage series mechanism can be represented by the following scheme: E3P
E3
E3S
E2P
E2
E2S
Tk~(1-n,p)Tk,
kcat'
9 E2+P+Q
Tk,(1-n,s) ~t
*- E I + P + Q
Kp K E: lactase; S: lactose; P: galactose (P); Q: glucose. Considering a f'mal inactive stage, the following expression is obtained:
":[I+A. k,
eo
k,
k2 - k I
(1)
k 2 -k I
Based on the modulation hypothesis, different first-order transition rate constants will exist for each enzyme species (free enzyme E, and secondary complexes ES and EP, equation (1) being applicable to any of them, with: k u = k i (1 - nij) (2) where subscript i refers to the inactivation stage, subscript J denotes the modulator and nu are modulation factors. In addition, the rate equation for lactose hydrolysis with CIL is [4]:
o,x, ....... l kcat "e v(X)
/ X + / m / +1 0 - x)
Km K;-1
K
so
From a material balance of all enzyme species, considering the proposed two-stage series mechanism, and the rate equation for lactose hydrolysis, enzyme inactivation under reactive conditions will be described by: de
= e - k 1 9[
dt
(1 - A). exp (- k I 9t)-[1 - a(X)-N I(X)] exp(- k 1 9t)+ (k I 9A (it 2 - k l ) )- [exp (- k , . t ) - exp (- k 2 9t)] +
A?k2"[exp(-k!'t),exP(,k2"t)]?[!-a(X)'Nz(X)]
]
+ (k 2 - k , ) e x p (-k, . t ) + k I 9A. [exp ( - k , . t ) - e x p ( - k 2 .t)]] where:
(4)
29 NI(X ) : n l s +nip .K m 9X / [ K p ( I - X ) ] "
N2(X ) =n2s +n2p . g m . X / [ K p ( I - X ) ]
2.2. Reactor performance under thermal inactivation Batch reactor performance will be described by the simultaneous resolution of equation (5), which represents the material balance over the reactor [6], and equation (4)" dX _ kc, , 9e(t).a(X) (5) dt so Packed-bed reactor performance, considering pseudo-steady-state and plug flow regime, will be described by the simultaneous resolution of equations (6), which represents the material balance over the reactor [6], and equation (4): dX e ( t , z ) . c r ( X ) - e - S
dz
F-s o
(6) 2.3. Temperature optimization Temperature has two opposite effects, increasing both reaction and inactivation rates. Kinetic and inactivation parameters are functions of temperature; therefore, reactor temperature can be optimized provided explicit functions for all of them are validated and a sound objective function is defined. Arrhenius-type functions can be proposed for all rate constants: kc, t =kclt,0 9e x p [ ' E T ]
;
Vmx=Vmx,0-exP[RET]
kij = ku'~ exp R . T ]
Affinity parameters (Km and Kp) can be considered as equilibrium constants and, therefore, thermodynamic correlations can be proposed: K
m
= K~0.exp R'T
;
K p = Kp, 0 - e x p AH~
LR.T
Such expressions can be introduced in equations (4), (5) and (6) to derive temperature explicit equations that will optimize reactor temperature. Specific productivity (the amount of product produced per unit mass of enzyme and unit time) may be used as a preliminary objective function. A more elaborated cost function will be the ultimate to consider.
3. METHODOLOGY 3.1. Materials Enzyme was a partially purified lactase (I3-D galactoside galactohydrolase, E.C.3.2.1.23) from Kluyveromyces marxianus var marxianus NRLL u produced and immobilized on activated chitin as already described [4]. A more stable CIL was used for temperature optimization studies, obtained by optimizing immobilization in terms of enzyme and glutaraldehyde to chitin ratios and particle size and including an acid pretreatment step. USP lactose was used for reactor runs. All other reagents were analytical grade fro m Merck (Darmstadt, Germany) or Sigma (Saint Louis, Mo, USA). One international unit (IU) of CIL
30 activity was defined as the amount of enzyme that hydrolyses one l~aol of lactose per minute at pH 6.6 and 40 ~ C in a 200 g/L solution. 3.2. Kinetics of enzyme inactivation Experiments in the absence and presence of modulators were done in batch reactors with temperature control. Air space was kept to a minimum to avoid enzyme inactivation. For experiments in the presence of lactose and galactose, medium replacement was done every eight hours. Lactose and galactose concentrations were 10"Kmand 10"Kl, to ensure equilibrium displacement to the corresponding enzyme complexes. In the former case, conditions were established to keep the enzyme saturated with substrate and enzyme to substrate ratio was low enough to keep conversion below 1%. Inactivation parameters were detennined by non-linear regression (Statistica, Stat Soft Inc, Tulsa, OK, USA) of experimental data to equation (1); modulation factors were determined from equation (2). 3.3. Reactor modelling and packed-bed reactor operation Model for reactor operation was obtained by solving equations [4] and [6] (or [5]), using modified Euler method. Computer resolution was done in Visual Basic 3.0 for Windows. Model was experimentally tested in a continuous packed-bed reactor with CIL. Reactor was a jacketed Pyrex-glass column, 2 cm in internal diameter and 18 cm in height, with catalyst bed height of 16 cm. Pseudo-steady-state was obtained after four to five residence times. 3.4. Temperature optimization Kinetic parameters at different temperatures were determined by linear regression of initial rate data in Lineweaver-Burk plots. Concentration range was 30 to 585 mM for lactose and for 30 to 180 mM for galactose. Inactivation parameters at different temperatures, in the absence of modulators, were determined as previously described. Temperature ranged from 15 to 40 ~ Temperature correlations for affinity parameters and rate constants were validated in Arrhenius-type plots.
4. RESULTS AND DISCUSSION 4.1. Kinetics of enzyme inactivation Kinetics of CIL inactivation at 28 ~ in the absence of modulators and in the presence of saturating concentrations of lactose and galactose is presented in Figure 1. Results were well represented by a two-stage series type mechanism of inactivation for all enzyme species, being a much better fit than one-stage first-order kinetics. From equation (1), shown by solid lines in Figure 2, inactivation parameters for the free enzyme E and the complexes ES and EP were determined by non-linear regression, as shown in Table 1. G-alactose was a positive modulator, modulation factors being positive in both stages. Lactase was a negative modulator in both stages of CIL inactivation. There are no reports on the particular effect of substrate and products on yeast lactase inactivation at operation temperatures. However, a two-stage mechanism of inactivation was proposed for the inactivation of yeast lactase in the presence of milk [7]. Experiments, however, were conducted at variable and non-saturating concentrations of sugars, obscuring conclusions. Negative modulation by substrate has been reported for glucose isomerase [8] and attributed to quaternary structure dissociation promoted by the substrate, which might be the case for lactose in K.marxianus lactase. Product modulation has
31 seldom been reported: in the case of penicillin acylase, strong protection was exerted by 6aminopenicillanic acid and negative modulation by phenylacetic acid [3]. Product protection can be explained by increasing hydrophobicity in the active site and by favouring a rigid configuration [9]. 100-
!. h
X X g
9
9
j.J
2O
0 0
|
!
5
i0
,
L
i
|
1.5
2O
30
35
40
415
TIE (Imam)
Figure 1. Kinetics of thermal inactivation of CIL at 28 ~ (X): phosphate buffer; (O): 545 mM lactose in buffer; (A): 870 mM galactose in buffer. Table 1 Inactivation parameters for CIL at 28 ~ according to a two-stage series-type mechanism Condition, Enzyme Specie Buffer E + galactose EP + lactose ES i i,
A
nlj
g~
k]. 102 (h'!) 7.74
k2"102 (hl ) 7.75
0.77
6.79
2.56
0.77
0.123
0.669
0.992
8.27
10.97
0.77
-0.068
-0.416
0.921
n2j
0.974
l, .,..
4.2. Reactor modelling and packed-bed reactor operation Packed-bed reactor operation was designed for an initial substrate conversion of 0.9. Reactor performance is presented in Figure 2. As seen, the model describes reactor performance very well and only in the first part conversion was somewhat higher than predicted. At prolonged times the model departs somewhat, but this is out of the region in which the inactivation mechanism was validated and is of no practical significance. Model not considering modulation factors was a very poor representation of reactor performance, which validates modulation factors as important parameters for proper reactor design. Enzyme decay profiles through catalyst bed height (data not shown) were consistent with the model, inactivation rate being lower near reactor outlet where protecting galactose concentration was higher.
32
0.$ o.B O,T
9
O.II
O.a i OA 0.3 0.2
9
0,1 0
.
"
l
0
10
20
! ....
30
40
Time (houri)
Figure 2. Continuous packed-bed reactor performance with CIL at 28 ~ a) model considering modulation; b) model not considering modulation; (e) experimental data. 4.3. Temperature optimization
Kinetics of CIL inactivation at different temperatures in the absence of modulators is presented in Figure 3. Results were in all cases well represented by a two-stage series-type mechanism of inactivation. As seen from figures 2 and 1, this CIL was more stable than the used formerly. Values for kinetic and inactivation parameters in the absence of modulators, at different temperatures, are presented in Table 2. Table 2 Kinetic and inactivation parameters of CIL at different temperatures Temperature (~ Km(m Vmax * 116.1 18.3 15 164.6 21.9 20 22.5 186.6 23.9 25 211.0 24.8 27.5 241.8 27.5 ....30 ..... 3il.7 31.7 .... 32.5 371.0 32.6 35 383.0 37.9 37.5 415.0 38.3 * ttmoles glucose/min.g catalyst ,
....
.
.
.
.
.
Kp (raM) 37.0 56.1 69.5 80.1
k]. 102 (h "l) k2.102 (h"z)
109.7 135.1
7.85 8.05 15.20 21.19
0.29 1.61 2.02 2.99
200.8
26.76
4.73
A
.
.
0.75 0.72 0.71 0.31 .
.
.
.
.
0.30
....
Thermodynamic correlations for temperature dependence were validated for Km and Kp. Arrhenius-type correlations were validated for kcat (Vmax)and ki, the following expressions being obtained: K m = 1.92" 10 .6 .exp R - T J
K p = 3.76-
.exp R . T J
33
Vmx
kI
--
fJ
F- 10.423
9.59.109
= 2.50-1013-
exp
I"-R--qr21"16]
k2
= 5.73- 10u
.I-" 38.20]
Correlation coefficients were in all cases around 0.99. A was not a defmite function of temperature, being rather constant at temperatures over and below 30 ~ However different values were obtained for both ranges: A = 0.73 T < 30 A = 0.31 T > 30 Batch reactor performance, not considering modulation effects (all n i j = 0), was simulated to determine optimum temperature, by solving equations (4) and (5) with temperature explicit functions for kinetic and inactivation parameters. Optimum temperature was determined considering specific productivity as objective function. Results are presented in Figure 4, temperature optimum being 26 ~ Kinetics on CIL inactivation in the presence of saturating concentrations of lactose and galactose is now underway to determine temperature explicit functions for modulation factors.
Figure 3. Kinetics of CIL inactivation in the absence of modulators:m: 25 ~ 27.5 ~ A: 30 ~ 4,: 32.5 ~ O: 37 ~
Figure 4. Temperature optimization for batch reactor with CIL
5. CONCLUSIONS .
CIL thermal inactivation was well described by a model based on a two-stage series mechanism, lactose and galactose being modulators. 9 A model for continuous packed-bed reactor performance with CIL, based on modulated thermal inactivation, was experimentally validated. 9 Explicit temperature functions were validated for all kinetic parameters and inactivation parameters in the absence of modulators
34 Batch reactor operation with CIL, not considering modulation effects, was simulated to determine optimum temperature. Results are underway to determine temperature explicit functions for modulation factors, to be included in the optimization model
REFERENCES 1. A. IUanes, M.E. Zufdga, Contreras, S. and A. Guerrero, Bioproc. Eng., 7 (1992) 199. 2. J.Y. Houng, H.Y. Yu and K.C. Chen. Biotechnol. Bioeng., 41 (1993) 451. 3. A. Illanes, C. Altamirano, M.E. Zufiiga, Bioteclmol. Bioeng., 50 (1996) 609. 4. A. Illanes, C. Altamirano, A. Aillap~n, G. Tomasello and M.E. Zuffiga, Enzyme Microb. Teclmol. (1998) (accepted for publication). 5. A. Sadana and J.P.Henley, Enzyme Microb. Technol. 7 (1985) 50. 6. E. CJalindo, O.T. Ramirez (eds.), Advances in Bioprocess Engineering, pp 467-472, Kluwer Academic Publ., Dordrecht, 1994. 7. R. Mahoney and T. Wilder. J. Dairy Res., 55 (1988) 423. 8. W. van den Tweel, A. Harder and R. Buitelaar (eds.), Stability and Stabilization of Enzymes, pp 111-131, Elsevier, Amsterdam, 1993. 9. I. Villaume, D.Thomas and M. Legoy. Enzyme Microb. Technol. 12 (1990) 506.
Nomenclature A molar activity ratio between initial and intermediate stage e enzyme activity e0 initial enzyme activity F actose feed flow-rate k~t catalytic rate constant k~ first-order transition rate constants Km Michaelis constant for lactose Kp inhibition constant by galactose R universal gas constant R 2 determination coefficient s lactose concentration so initial (inlet) lactose concentration S cross section of reactor t time T temperature v reaction rate Vma~maximum reaction rate ( = l~t. e) X substrate conversion ( = [s0-s]/s0) z axial dimension in reactor s liquid volume fraction in catalyst bed
Acknowledgements This work was supported by Grant 1971029 from FONDECYT, Chile
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
35
Fast in-situ-characterisation o f b i o c a t a l y s t s l o n g - t e r m stability M. Boy, A. Dominik, H. Voss SFB Biocatalysis, Institute of Biotechnology, Graz University of Technology, Petersgasse 12, A-8010 Graz, Austria
1. INTRODUCTION A major bottleneck in developing industrial biocatalysts is the lack of a method to determine the catalyst process stability quickly and efficiently [1]. In particular, the determination of these properties is difficult, often inaccurate and expensive because highly active, selective and stable biocatalysts should be achieved [2,3,4]. To solve this problem, a new method for fast in-situ-characterisation of activity and longterm stability of enzyme carrier catalysts has been developed. To accelerate the process of ageing the temperature is increased continuously during the experiment. On the basis of the proposed dynamic model it is possible to determine information pertaining to enzyme kinetics, selectivity, interaction between carrier, enzyme and substrate as well as information regarding the stability of the enzyme carrier complex, including reversible and irreversible activation and inactivation parameters. In contrast to common, isothermal techniques these parameters can be estimated for the whole temperature range in one short experiment. Working under industrial process conditions allows one to determine the whole relevant information necessary to characterise the (carrier- bound) biocatalyst. This knowledge allows the design and optimisation ofbiocatalytic industrial processes within a short period of time [5,6]. 2. T H E O R Y
To demonstrate the method of fast in-situ-characterisation of activity and long-term stability of enzyme carrier catalysts a single step deactivation with reversible deactivation of the native and active enzyme F_~ is presented. The equilibrium constant K gives the ratio of reversible deactivated enzyme ED. and F_~. The sum of Es and ED, deactivates irreversibly to ED with the deactivation rate coefficient k d.
~E~* In mathematical terms the mechanism can be described by a mass balance and a differential equation:
[Eo,]+[E,]
(1)
36
.[e]
dt
At the beginning of the process the whole enzyme is in the native form. The deactivation rate coefficient kd and the equilibrium constant K are temperature dependent according to the Arrhenius law: Eo
ka = ka.| -e R.r
(3)
[E~
(4)
[E I Ed.r,,v
K=K|
R.r (5) In addition to the reversible and irreversible enzyme deactivation, the biochemical reaction is also a function of temperature. In general, the biochemical reaction rate can be expressed in separable terms of temperature T, substrate concentrations ci and enzyme concentration Cen~me(eq. (6)). A frequently used expression for the kinetics of enzymatic reactions is the Michaelis-Menten-equation. It is possible to define the maximum reaction rate rm~x as a product of rate coefficient for the enzyme-catalysed reaction k~ and the native enzyme concentration [Es], as demonstrated in equation (7). In the case of substrate saturation equation (7) simplifies to (8). Adequate substrate concentrations and flow rates were chosen during the experiments to fulfil this assumption.
r= f ( r , ci,ce~m)= f, (r). f2 (ci). f3 (Cenzym )
IS] "K. [S--------] + __kr
r-r
.[EN ]
(6) (7)
The definition of the specific enzymatic activity "a" [mol/g.s] in equation (9) and (8) together with (10) for the temperature dependence of the rate coefficient k~ finally leads to equation (11): (9) a--~rmax
[E]o
k r = h| "e R.T = A
(10)
[E~] -e-L [EN] a = A. [El0' = A.~-e "'r 9[El0
(11)
The set of equations (1)-(5) and (11) allows the calculation of the time course of the specific enzymatic activity dependent on temperature. 3. MATERIALS AND METHDOS 3.1. Enzymes and chemicals The goal of the present study is to demonstrate the application of the newly developed method for characterising free and carrier-bound enzymes during the development of carrierbound biocatalysts. A lipase of Mucor miehei was examined both as a free enzyme and immobilised on different carriers. Novozyme 388 (Novo Nordisk) was used as a free enzyme. Additionally, a carrier bound enzyme of this source is commercial available from Novo Nordisk (Lipozyme IM). Support material is a highly porous anion exchanger resin of the
37 phenolic type (particle size 0,2-0,6mm). As a parallel enzyme, we immobilised Novozyme 388 adsorptive on a hydrophobic organopolysiloxane-carrier (DELOXAN ~, propylsiloxane modified ethylenediaminopolysiloxane, particle size 0,2-0,4 mm, Degussa AG). The carrier was saturated with 170 ~gprotein/gdry carrier with a yield of 80 %. A highly active carrier-bound enzyme with 30000 IU/g~y c~meraccording to the tributyrine standard assay [7] is achieved. As a model reaction, the hydrolysis of triglycerides was chosen. Tributyrine (Fluka) was emulsified in water according to the literature[7]. 3.2. Experimental conditions
Experiments with the free enzyme were carried out in jacket-cooled, flat-flange standard glass reaction vessels with 2000 ml total volume (Schott). A fed-batch process was used. The initial reaction volume was 200 ml. The feed rate depends on the enzymatic activity. Substrate saturation should be achieved during the whole experiment. The feed rate to fulfil this assumption, the used enzyme amount and the applied temperature program are presented in Table 1. The feed rate was attained by a gear pump (MC-Z, Ismatec). The substrate solution was stored under stirring on a magnetic stirrer in a ice bath. The speed of a 3-blade-impeller stirrer with 70 mm diameter was 300 rpm. Temperature control was reached by a computercontrolled external heating bath. pH-value was kept constant at pH 8 using an autotitrator (Schott Titroline). We used 0,1 mol/1 potassium hydroxide solution as a titrant. For immobilised enzymes continuous processes in a jacket-cooled, fiat-flange standard glass reaction vessel with 500 ml total volume (Schott) and 250 ml working volume were carried out. 300 rpm stirrer speed of the used 3-blade-impeller stirrer with 70 mm diameter was found to be enough to neglect external mass transfer resistance. The flow rate depends both on enzymatic activity and the amount of enzyme amount used. Flow rates, enzyme amount and temperature programs are presented in Table 1. The feed was attained by a gear pump (MC-Z, Ismatec). The working volume was held constant by pumping out the excess solution by a vacuum filtration aspirator pump through a glass frit. Due to higher activity, 1 mol/1 potassium hydroxide solution was used as titrant to keep the pH-value constant at pH 8. The time course of titrant necessary to neutralise the fatty acids produced is automatically recorded, corrected and converted into activity. Table 1 Experimenta! conditions Carrier Reactor Feed rate Enzyme Enzyme Enzyme material type [ml/h] amount preparation source fed-batch 200 14 pl Free enzyme Novozyme 388 528 0,015g ~ = Immobilised Novozyme organopoly- continuous siloxane stirredtank enzyme 388 Immobilised LipozymeIM enzyme
anion
reactor continuous
resin
reactor
exchanger stirredtank
214
0,499gd~~
Temperature Program 10-70 ~ in 7 h const. 6,4 ~ for 2 h, 6,4-66,4 in 7h 10-70 ~ in 7 h
38 4. PARAMETER ESTIMATION x
measured
,,
c a l c u l a t e d specific activity
........
native e n z y m e c o n c e n t r a t i o n
..... 25000
,
'
temperature ,
,
9
. . . . . . . . . . . . . . . . . . . . . . . 9 .-
E" .v
specific activity
20000.
.
,,
i ~
, ~
,
,
-
s
"-.
0,03
x
10000,
o,o2
O
5000
O
0
(D
0,05 ~ 0,04~
~
15000,
~r
0,06 %; 70
l !
s
~ > .n
|
0
~
~
~
.
.
~ .
0,00
10
O 50 .o .
-~-
40
c:
30~
8 8
0,01 E .
6 0 ,..., == =
"
|
20E
10F-
N
c "'
Time [h]
Figure 1. Cleavage of tributyrine by Novozyme 388 immobilised on organopolysiloxane. Figure 1 shows the measured and simulated specific activity, the temperature profile and the computed concentration of native enzyme of the lipase Novozyme 388 immobilised on organopolysiloxane. In order to determine the temperature dependent coefficients of equations (3), (5) and (11), the measured specific activity is evaluated in sections. In the first time interval ( 2 - 4 . 5 h) it is possible to neglect reversible and irreversible deactivation of the enzyme because of the low temperature (6-26~ and short operation time. This assumption leads to equation (12). If the enzyme is saturated by a high substrate concentration or by a high flow rate, equation (11) finally simplifies to (13): [EN]/[E]o=I.
(12)
E.
a = A| .e R.r (13) By presenting the specific activity during this time interval in an Arrhenius plot, the activation coefficients A~ and E=could be determined. With increasing temperature the fraction of reversibly deactivated enzyme increases. Irreversible deactivation plays a minor role because of moderate temperature and operation time. A mass balance of the enzyme leads to equation (14). Combining equation (4) and (14) with (11 ) results in equation (15) to determine the equilibrium constant K:
(,4)
[E]o = _ E....~a
K
R.r -1 (15) a During the time interval between 5 and 6.2 h the equilibrium constant K could be calculated from the measured specific activity "a" and from the previously determined activation coefficients A~ and E,. An Arrhenius plot of K leads to the reversible deactivation coefficients K~ and Er At the end of the temperature program, the fraction of irreversibly denaturated enzyme increases. When combining equations (1), (2), (4) and (11), one finally obtains a formula allowing an evaluation of the coefficients of irreversible deactivation. =
A|
39
ka
=
/
K
I+K
.
Ear~ , _ R
.
d 1 T___.~I dt a
da dt
(16)
The equilibrium constant K is temperature dependent according to equation (5). During the time interval between 7 and 8 h k d, could be calculated from the measured specific activity "a" and the previously determined activation and reversible deactivation coefficients. The graphical presentation of kd in an Arrhenius plot leads to the determination of the irreversible deactivation coefficients k~| and Ea, irr.
5. RESULTS AND DISCUSSION The presented method was used to determine the influence of immobilisation on the examined lipase. For all experiments carried out to determine the influence of immobilisation, a first-order deactivation mechanism with a reversible deactivation was found to be the better fitting compared to a first-order deactivation mechanism without reversible deactivation. It has to be stressed in general, that it is not the claim of the presented method to proof a mechanism, but to find a model with a set of parameters which fits the experimental data well and can be used to calculate enzyme characteristics and process strategies. The coefficients of the different immobilised and free enzymes are presented in Table 2. Half-life time (Fig. 2) and "Turn-over-Number" (TON) (Fig. 3) were calculated. TON expresses how much substrate can be converted until a defined residual activity is reached.
TON =
~a(t)dt (17) 0 For example, the residual activity can be given by a minimum conversion with a fixed amount of enzyme used in a reactor. Table 2 Activation and deactivation coefficients of free and carrier-bound M u c o r m i h e i lipase. Activation coefficients Deactivation coefficients Reversible Irreversible A| E, K,o E~ k,t| E~i, [mol/g.s] [kJ/mol] [.] [kJ/mol] [I/s] [kJ/mol] Novozyme388 175,5 32,53 3,6561019 120,52 5,2311033 231,79 Novozyme 388/ 13,45 26,74 7,5671021 134,44 6,733.103s 262,66 organopolysiloxane Lipozyme IM 16~5104 60,19 1,3631019 114~50 1,6171049 328,10 Figure 2 clearly shows the big advantage of the presented method for fast in-situcharacterisation of activity and long-term stability. The half-life time ofLipozyme IM is about 140 days. This period is necessary to measure the half-life time in an isothermal experiment. To determine the half-life times of Lipozyme IM in the whole temperature range from 10 to 70 ~ in 10~ intervals using the common isothermal techniques would take about 3700 years (of course, even during 140 days other phenomena beside thermal deactivation (e.g. microbial growth) may be responsible for shorter half-life times). To get the same information using the new method one 12 h-experiment is enough. Figure 3 shows the TON of free and immobilised M u c o r m i h e i lipase. 300 U/g were defined as minimum specific activity. Lipozyme IM reaches this lower level only in the
40 temperature range between 28 and 54~ It has a maximum in the temperature course of the TON. This temperature is an optimum operation point if the costs of the enzyme plays an important role in the overall process. 101~
Lipozyme IM [ ....... Novozyme 388
108 ~" 106 11)
E
....,
"', ,",,"-',-',"~='~"~~~...,..~."
o
E
388
1016
101'
,.:..
-~- 101z Z O 101~ t" 108 "~ Os
104 102
~_. 100
Lipozyme IM ....... Novozyme 388 ~:.~:. :... immob. Novozyme 388 ...,.
~":':.-..
0 1 E =.1 04
"r" 10 .2
lo
2o
3o
40
so
60
A
a0
Temperature [*C] Figure 2. Half-life time of free and carrierbound Mucor mihei lipase.
Z 102
0
I--
:.. 9.
9
0
,
,
,
.
,
.
,
.
,
.
|
,
!
,,.,
,
"'|'.."
10 20 30 40 50 60 70 80 90
Temperature [*C] Figure 3. Turn-over-Number of free and cartier-bound Mucor mihei lipase.
6. CONCLUSIONS The new developed method for fast in-situ-characterisation of activity and long-term stability of enzyme carder catalysts enables researchers, producers and users of enzymes and carrier-bound enzymes to characterise biocatalysts quickly and, therefore, accelerates the process of choosing and developing biocatalysts and carriers. Commercial available biocatalysts could be very easy characterised for a special problem. A quick process optimisation regarding process conditions, reaction media and finally the costs of the catalyst thus becomes possible. REFERENCES
1. Cabral, J.M.S., Best, D., Boron, L., Tramper, J.: Applied Biocatalysis, Harwood publ., Chur 1994. 2. Mozhaev, V.V.: Mechanism-based strategies for protein thermostabilization. TIBTECH Vol. 11, 1993. 3. Gupta, M.N. (ed.): Thermostability of enzymes. Springer, Berlin 1993. 4. Vrabel, P., Polakovic, M., Stefuca, V., Bales, V.: Analysis of mechanism and kinetics of thermal inactivation of enzymes: Evaluation of multitemperature data applied to inactivation of yeast invertase. Enzyme and Microbial Technology 20, 348-354, 1997. 5. VoB, H.: DP 195 46 192.4 1995. 6. Boy, M., Dominik, A., Vog, H.: Fast determination of biocatalysts process stability. Process Biochemistry (submitted). 7. Weber, K. H., Stecher, H., Faber, K.: Some Properties of Commercially Available Crude Lipase Preparations. In: Preparitive Biotransformations, Editor: S.M. Roberts, Wiley, New York, 1995, S. 5:2.1-5:2.10.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Chemical
and t h e r m a l
stability
41
of ferulic
acid (feruloyl)
esterases
from
AspergiUus. C.B. Faulds ~, F.O. Aiiwan ~, R.P. de Vries b, Williamson ~.
ILW. Pickersgill ~, J. Visser b and G.
aBiochemistry Department, Institute of Food Research, Norwich Research Park, Colney, Norwich NR4 7UA, UK. bMGIM, Wageningen Agricultural University, Dreijenlaan 2, 6702 HA, Wageningen, The Netherlands CFood Macromolecular Science Department, Institute of Food Research, Early Gate, Reading RG6 6BZ, UK. ABSTRACT Ferulic acid (feruloyl) esterases are a novel subclass of the carboxylic acid esterases, and are able to hydrolyze esterified phenolic acids from plant cell walls. The active sites of feruloyl esterases show high homology with the serine active sites of lipases, and a model of a feruloyl esterase (FAE-III) from Aspergillus niger was derived based on the structure of four lipases. Stability of this feruloyl esterase was compared to a feruloyl esterase from A. tubingensis using chemical and thermal denaturation. These enzymes show a 92.5% identity in their amino acid sequences. The A. tubingensis feruloyl esterase is more sensitive to degradation than the A. niger FAE-III. These enzymes have potential in the food and agro-chemical industries.
1. INTRODUCTION Ferulic acid (feruloyl) esterases (FAE) are a subclass of the carboxylic acid esterases (EC 3.1.1.1) and are able to hydrolyze esterified phenolic acid from plant cell-walls. These types of linkages are found in a wide variety of cell-walls [1 ], and the enzymes are secreted by the microorganism to facilitate total breakdown of the cell-wall polymers. Feruloyl esterases have been purified from bacterial and fungal sources [e.g. 2-8], and are induced during growth on lignocellulosic material. They differ from other cell-wall degrading esterases in their specificity for cinnamoyl esters (as shown by Km values between 10-6 and 2 x 10.5 M for some substrates). Five genes encoding enzymes with feruloyl esterase activity have recently been sequenced [5, 810], showing very little conserved identity, apart from the amino acids adjacent to the active site serine. Modification of nucleophilic serine residues has indicated a catalytic role for serine in these esterases [4], and the GXSXG motif common in lipases together with aspartate and histidine residues which make up the catalytic triad appear to be conserved. From its primary sequence, a model of the tertiary structure of FAE-III from Aspergillus niger was based on
42 comparisons with lipases, in open and closed lid conformations, from Rhizopus delemar, Rhizomucor miehei, Humicola lanuginosa and Penicillium camembertii ([10], Aliwan, Faulds, Pickersgill & Williamson, unpublished results). Aspergillus species have been shown to produce a number of feruloyl esterases, but the gene encoding one (faeA) has been sequenced to date from two species, A. niger and A. tubingensis [ 10]. This gene shows a high degree of nucleotide sequence identity (87.1%) between the two strains and contains one conserved intron. Translation of the DNA sequence of both genes resulted in amino acid sequences of 281 and 280 for A. niger and A. tubingensis, respectively, each containing a conserved signal peptide of 21 amino acids. The esterases from A. niger (FAE-III) and A. tubingensis contain a single putative N-glycosylation site, and an alignment of their amino acid sequence showed a 92.5% identity, with a calculated molecular mass of 28314 and 28261, respectively [ 10]. In this report, we compare the A. niger and A. tubingensis gene products over-expressed in a protease-deficient strain of A. niger and subsequently purified, and discuss possible reasons for one form to be more resistant to proteolysis. 2. MATERIALS AND METHODS Fungal strains and culturing. A. niger NW154 containing a prtF mutation (pyrA6 prtF28) and A. tubingensis NW241 (pyrA2 fwnA1) have been described previously [10]. Functional constructs containing the faeA genes were transformed into either the A. niger NW154 prtF mutant or A. tubingensis NW241 [10]. Purified esterase was obtained from A. niger NW154 transformants. Cultures were grown on 1% (w/v) oat spelt xylan containing minimal media for up to 6 days at 25~ Supernatants were collected by filtering through miracloth. Purification and characterization of feruloyl esterases. Recombinant esterase was purified from culture supernatants after 3 days of growth on oat spelt xylan as described previously [3]. Molecular masses were determined by SDS-PAGE (Novex 10% Tricine gels) calibrated with known standards and by electrospray mass spectroscopy on a Micromass Platform positive spray mass spectrophotometer. Western analysis of culture supernatant samples was performed using polyclonal antibodies raised in mice against purified FAE-III. Enzyme assays:. Feruloyl esterase activity in culture supernatants and throughout the purification was determined using methyl ferulate (50 lxM) as a substrate at pH 6.0 and 37~ [3]. One unit of enzyme activity is the amount of enzyme which released 1 lxmol of product per minute under these conditions. Chemical and thermal inactivation of FAE-III were Chemical and thermal inactivation. determined as previously described [ 11]. Determination of N-terminal sequences. Amino acid sequences were determined from electroblots of an SDS-PAGE separation of purified A. tubingensis FAE on an Applied Biosystems Procise Sequencer. Modeling of FAE-III. The A. niger FAE-III model was generated using the program MODELLER [ 12], with four aligned lipases giving the input protein scaffold and the sequence alignment giving the relationship between this scaffold and the A. niger FAE-III sequence. The
43 molecules were displayed on a Silicon Graphics Indigo 2 with the program O [13] and the protein cartoon was generated using MOLSCRIPT [ 14]. 3. RESULTS AND DISCUSSION Over-expressing transformants of both A. niger and A. tubingensis, containing either thefaeA gene from A. niger or the faeA gene from A. tubingensis, were grown for 6 days, culture supernatants concentrated 5-fold and subjected to Western analysis (Figure 1). On SDSPAGE, multiple bands were visible (results not shown), ranging from the mature protein at 36 kDa (approx. 30% of total protein for the A. tubingensis FAE and 95% for the A. niger FAEIII) to smaller bands down to 22 kDa. Upon Western blot analysis, it was apparent that only the A. tubingensis enzyme gave a multiple banding pattern, irrespective of the host strain of Aspergillus. The recombinant esterase from A. niger (FAE-III) showed partial degradation when expressed in A. tubingensis, only upon Western analysis (Figure 1, lanes 1 and 3). Thus, it appears that the A. niger feruloyl esterase is less sensitive towards degradation by natural Aspergilli proteases.
Figure 1. Western blot degradation patterns for feruloyl esterases from Aspergillus. Lane 1, transformant NW154::pIM3207.7 (A. niger faeA in A. niger); lane 2, NW154::pIM3208.5 (A. tubingensis faeA in A. nige0; lane 3, NW241::pIM3207.7 (A. niger faeA in A. tubingensis); lane 4, NW24 l::pIM3208.5 (A. tubingensis faeA in A. tubingensis); lane 5, purified FAE-III from A. niger [3]. Cultures grown on oat spelt xylan for 6 days. With courtesy of Appl. Environ. Microbiol. [ 10]. The recombinant esterase from A. tubingensis was purified by ammonium sulphate precipitation, hydrophobic interaction chromatography and anion-exchange chromatography. SDS-PAGE of each stage of the purification still resulted in the presence of multiple bands for the denatured enzyme. Mass spectroscopy of the enzyme aider anion-exchange revealed a single mass of 29724, compared to the A. niger FAE-III mass of 29756. Thus, the feruloyl esterase from A. tubingensis was apparently cleaved at certain points on the protein, but this did not appear to alter the native molecular mass. From Figure 1, there appears to be a number of these cleavage sites. N-terminal sequencing of the first five residues of the mature A. tubingensis protein (36 kDa) and the second largest band on SDS-PAGE (26 kDa) gave identical results, presumably the lower band is produced by C-terminal truncation of the mature protein. A well separated band at 22 kDa was also N-terminally sequenced (5 residues). This band contained two sequences of equal signal: one corresponding to the N-terminal sequence (A S T Q G); the other a sequence X
44 E V H, which corresponds to a sequence 95 residues in from the N-terminus (Figure 2). In the protein sequence for the A. tubingensis esterase, X corresponds to cysteine. Figure 2 shows an alignment of the two Aspergillus feruloyl esterase sequences. There are 15 amino acid changes between the two sequences, and one extra residue at position 185. The degraded band sequence at 22 kDa encompasses one of these amino acid changes at the point of cleavage: A. niger (D *C E V H) and A. tubingensis (S *C E V H). Thus, it appears that an aspartic acid residue instead of a serine at position 93 may contribute to the proteolytic stability of FAE-III. ANFAE ATFAE
ASTQC ASTQC
ANFAE ATFAE
ISEDL YNRLV ISEDL YSRLV
EMATI EMATI
SQAAYADLCN SQAAY ADLCN
IYNAQ IYNSQ
45 45
TDING WILRD DTSKE TDING WILRD DSSKE
IITVF RGTGS DTNLQ LDTNY TLTPF DTLPQ IITVF RGTGS DTNLQ LDTNY TLTPF DTLPQ
90 90
ANFAE ATFAE
CNDCE VHGGY YIGWI CNSCE VHGGY YIGWI
S V Q D Q V E S L V KQQAS Q Y P D Y A L T V T G H S L G SVQDQ VESLV QQQVS QFPDY / ~ ' r V T GHSLG
135 135
ANFAE ATFAE
ASMAALTAAQ ASLAA LTAAQ
ANFAE ATFAE
P E T T Q Y F R V T HSND_G IPNLP P A D E G Y A H G G V E Y W S P D T T Q Y F R V T HAND_G IPNLP P A D E G Y A H G V V E Y W S
ANFAE ATFAE
VCTGD EVQCC VCTGD EVQCC
LSATY DNVRL YTFGE LSATY DNIRL YTFGE
IPSTI IPSTI
PRSGN QAFAS PRS N Q A F A S
E A Q G G Q G V N D A H _ T T Y FGMTS E A Q G G Q G V N N AH_TTY FGMTS
GACTW GHCTW
IKGEK IKGEK
YMNDA YMNDA
FQVSS FQASS
180 179
VDPYS AQNTF VDPYS AQNTF
225 224 260 259
Figure 2. Alignment of the deduced amino acid sequence for mature feruloyl esterases from A. niger and A. tubingensis. The sequence identity is 92.5%. The region homologous to the putative serme lipase catalytic triad is underlined, and the points of change are indicated by a * (from [101)
A model of A. niger F AE-III (Figure 3), based on sequence identity with four fungal lipases, shows that the location of this amino acid change lies on an exposed loop of the ct/13 barrel structure, and so the residue is accessible to proteolytic attack while in the native form. Activity of the purified esterases from A. tubingensis was compared to FAE-III from A. niger against methyl ferulate. Activities were very similar:- FAE-III (4.38 l.tmol/min/mg protein) and A. tubingensis FAE (4.46 U/mg protein). This indicates that the differences in amino acids (16 residues) are not important for activity on methyl ferulate, and that proteolytic attack on A. tubingensis FAE does not interfere with the catalytic mechanism. Initial results on the stability of the two esterases to thermal inactivation suggests that they behaved in a very similar manner at 60~ losing 50% of their activity in 60-80 minutes. Thermal inactivation of FAE-III from A. niger has been found to be reversible, the first order rate constant of unfolding (v) was 0.76'10 .3 per second, and AG* was 101.9 kJ/mol [11].
45
Figure 3. A schematic ribbon diagram of the 3-D fold of A. niger FAE-HI monomer model. The model was based on the alignment of FAE-III with four lipases: Humicola lanuginosa [ 15], Penicillium camembertii [16], Rhizomucor miehei [17] and Rhizopus niveus [18], which showed significant homology (Z-score larger than 4.0). The N-terminus is located at the bottom right of the figure and the C-terminus is located near the active site. The location of the loop containing the D'-~S amino acid change is indicated. In summary, two feruloyl esterases from Aspergillus show 92.5% amino acid sequence identity, similar physical characteristics, and specific activity against methyl ferulate, but the enzyme from A. niger (FAE-III) is more resistant to degradation than the esterase from A. tubingensis.
This work was supported by a Biotechnology and Biological Sciences Research Council (BBSRC) Realizing Our Potential Award (ROPA), the Industrial Research Center (Tripoli, Libya) for a Ph.D. scholarship to FOA, and the BBSRC, UK. We wouM like to thank Dr. Fred Mellon (Institute of Food Research) for carrying out the electrospray mass spectroscopy on the two enzymes. REFERENCES
[1] [2] [3] [4]
R.D. C.B. C.B. P.A.
Hartley and E.C. Jones, Phytochemistry 16 (1977) 1531. Faulds and G. Williamson, J. Gen. Microbiol. 137 (1991) 2339. Faulds and G. Williamson, Microbiol. 140 (1994) 779. Kroon, C.B. Faulds and G. Williamson, Biotechnol. Appl. Biochem. 23 (1996) 255.
46
[5] [6]
[7] [8] [9] [10]
[11] [12] [13] [14] [15] [16] [17] [18]
L.M.A. Ferreira, T.M. Wood, G. Williamson, C. Faulds, G.P. Hazlewood, G.W. Black and H.J. Gilbert, Biochem. J. 294 (1993) 349. W.S. Borneman, L.G. Ljungdahl, R.D. Hartley and D.E. Akin, Appl. Environ. Microbiol. 58 (1992) 3762. A. Castanares, S.I. McCrae and T.M. Wood, Enzyme Microb. Technol. 14 (1992) 875. B.P. Dalrymple, Y. Swadling, D.H. Cybinski and G-P. Xue, FEMS Micro. Letts. 143 (1996) 115. B.P. Dalrymple and Y Swadling, Microbiol. 143 (1997) 1203. R.P. de Vries, B. Michelsen, C.H. Poulsen, P.A. Kroon, R.H.H. van den Heuvel, C.B. Faulds, G. Williamson, J.P.T.W. van den Hombergh and J. Visser, Appl. Environ. Microbiol. 63 (1997)4638. G. Williamson and J. Vallejo, Int. J. Biol. Macromol. 21 (1997) 163. A. Sali and T.L. Blundell, J. Mol. Biol. 234 (1993) 779. T.A. Jones, J-Y. Zou, S.W. Cowan and M. Kjeldgaard, Acta Cryst A47 (1991) 110. P. Kraulis, J. Appl. Cryst. 24 (1991) 946. U. Derewenda, L. Swenson, Y. Wei and Z.S. Derewenda, J. Lipid Res 35 (1994) 524. S. Yamaguchi, K. Takeuchi, T. Mase, K. Oikawa, T. McMullen, U. Derewenda, R.N. McElhaney, C.M. Kay and Z.S. Derewenda, Prot. Engin. 9 (1996) 789. Z.S. Derewenda, U. Derewenda and G.G. Dodson, J. Mol. Biol. 227 (1992) 818. M. Kohno, J. Funatsu, B. Mikami, W. Kugimiya, T. Matsuo and Y. Morita, J. Biochem. 120 (1996) 505.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
47
Stability Properties o f C a l f Intestinal Alkaline P h o s p h a t a s e Ultan McKEON, Brendan O'CONNOR & Ciarhn 0'F/kG.~qq" School of Biological Sciences, Dublin City University, Dublin 9, Ireland.
Inactivation of alkaline phosphatase (APase) in the range 48-65~ fits a first order decay; an Arrhenius plot gives estimates of low-temperature half-lives. APase tolerates urea but not GdC1. Micromolar amounts EDTA lead to inactivation, as do millimolar amounts 1,10phenanthroline. APase is sensitive to SDS but retains significant activity in presence of the solvents DMSO, DMF or THF (up to 60% (v/v) in buffer).
1. INTRODUCTION Alkaline phosphatase (Orthophosphate monoester phosphohydrolase; EC 3.1.3.1; APase) is used to prevent self-annealing of DNA by cleavage of 3'-terminal phosphate groups (1). Its ability to utilize luminescent substrates has led to its use as a reporter enzyme in nonradioactive DNA probes (2) and in sensitive enzyme immunoassays (3). Its levels in serum can be diagnostic in clinical chemistry (4) and it is included in commercial control sera. Long-term persistence of enzyme activity can be a critical parameter in these various types of diagnostic or molecular biology products. It would be useful to have some indication of the baseline stability characteristics of APase and of its ability to withstand various kinds of inactivating influences. The tetrameric, zinc-containing calf intestinal APase (5) is widely used. Here we explore the intrinsic stability properties and limitations of this enzyme and we report on its tolerances of heat, denaturants, chelating agents, detergent and water-miscible organic solvents.
2. EXPERIMENTAL Boehringer Mannheim calf intestinal APase (product # 108 138) was diluted 1/1500 in 2amino-2-methyl-1-propanol/metal ion buffer (6) for use and assayed by the method of IFCC [intemational Federation of Clinical Chemistry] (6). 4-nitrophenyl phosphate and other assay reagents were from Sigma, as were urea, guanidine chloride [GdC1], EDTA, 1,10phenanthroline and SDS. Aldrich supplied 1,7- and 4,7-phenanthrolines.
"Author for correspondence.Tel +35317045288; Fax +35317045412; Email
[email protected]
48 Dimethylsulphoxide [DMSO] was from BDH while acetone, dimethylformamide [DMF] and tetrahydrofuran [THF] were from Labsean (Dublin, Ireland). For the temperature profile, aliquots of APase were placed (after preheating to 35-40~ for 10 min at temperatures in the range 35-70~ then removed onto ice to prevent further denaturation. Residual activity was later determined under optimal conditions. APase was heated continuously at 48~ 50~ 55~ 60~ and 650C to obtain thermoinactivation curves. Residual activities of samples, removed at intervals and stored on ice, were assayed. An exponential decay curve of remaining activity against time was plotted to estimate the first order rate constant, k, at each temperature. Urea and GdC1 stock solutions (5M) were prepared according to (7), but using Tris, pH 8.3, instead of MOPS buffer. Incubation of the enzyme in the denaturant took place at 30~ for 2 h, after which time enzyme activity was determined. The effects of EDTA (pH 8.3), 1,7-phenanthroline, 1,10-phenanthroline and 4,7phenanthroline on activity were determined. After incubation at 370C for 4h, samples were extensively dialysed (40C) against 10mM Tris-HC1, pH 8.3. Activities were then measured. Enzyme was treated with various v/v concentrations of THF, DMF and DMSO and assayed after 90 min at room temperature. APase was treated with various w/v concentrations of SDS. After 2h at 30~ catalytic activity was measured.
3. RESULTS & DISCUSSION
Thermostability Studies. The temperature profile experiment gave 57~ as halfinactivation temperature, Tso. All thermoinactivation curves in the range 48~176 fitted a first order decay curve and gave linear semi-log replots. Values for the first-order rate constant, k, and half lives (t~t2= 0.693/k) are shown in Table 1. Table 1 Values of first-order rate constant, k, for APase thermoinactivation Temperature
48~
50~
55~
60~
65~
k, min"
0.021+ 0.001
0.028+ 0.001
0.063+ 0.002
0.143+ 0.008
0.30+ 0.04
An Arrhenius plot of these data yielded a value of 142 kJ.mol-! for the inactivating event under the conditions used. Extrapolation of the Arrhenius plot permits estimation of k (and of likely half lives) at temperatures of interest - see Table 2. Note that the value for 0~ takes no account of possible freezing effects. The values obtained under our particular experimental conditions may not be generally applicable. Franks (8) has discussed strengths and weaknesses of accelerated stability testing.
49 Table 2 APase half-lives estimated from Arrhenius plot. Temperature
0~
4~
20~
25~
37~
Half-life, h
6360
2568
88.8
33.6
3.6
Half-life, days
265
107
3.7
1.4
0.15
The DNA dephosphorylation protocol of Maniatis et al. (1) used calf intestinal APase for 1 h at 37~ followed by inactivation at 68~ for 15 min in SDS. The present results indicate that the enzyme remains active throughout the 37~ incubation and will quickly undergo inactivation at 68~ In the measurement of PCR products, Yang et al. (9) incubated APase at 37 ~ for 30 min in an enzyme-substrate reaction. Nakagami et al. (10) used APase in an enzyme-conjugated DNA probe system; they carried out hybridization of probe with target DNA at 42~ and found that a 30 min hybridization period gave the greatest sensitivity, with decreased sensitivities at longer times. These protocols appear to make good use of the intrinsic thermal stability characteristics of APase. de la Fourniere and colleagues (11) studied thermal and pH stabilities of bovine intestinal APase by Fourier transform infrared spectroscopy [FTIR]. At temperatures up to 70~ (or at pH values down to 5.4), activity loss was not accompanied by significant FTIR changes; i.e. there were no notable conformational alterations. At 80~ (or at pH 3.4), however, APase began to unfold after it had completely lost activity. FTIR results were confirmed by H/2H exchange and circular dichroism. Denaturant Stability. Fig. 1 shows the effects of increasing concentrations of urea and GdCI on APase activity. Urea up to 3M exerts a slight activation effect and approximately 80% activity is retained even at 5M urea. GdC1 has a similar activating effect up to 0.5M but APase activity declines steeply at GdC1 concentrations above 0.5M and is almost absent at 2M GdC1. GdC1 is a more potent denaturant than urea (7). At low concentrations, both denaturants slightly activate APase, perhaps by altering its conformation slightly to favour catalysis. 0.1M GdC1 caused tertiary structural changes in the monomeric horseradish peroxidase without affecting activity (12). Nakagami et al. (10) noted that APase retained 40% activity after 3 h in 6M urea. Effect of EDTA and Phenanthrolines. Micromolar EDTA concentrations have a potent adverse effect on APase (Fig. 2), as one would expect for a Zn-containing enzyme. This is most likely due to chelation of the catalytically-essential Zn 2§ ion and is confirmed by Fig. 3, which shows the effects of phenanthrolines on APase activity. The chelating 1,10phenanthroline adversely affects APase (but at millimolar concentrations) while the nonchelating 4,7- and 1,7-phenanthrolines (13) have little or no effect. Note that exhaustive dialysis of EDTA-containing samples took place prior to assay to prevent EDTA interference in the assay mix. Femley (5) has stated that the kinetics of EDTA inhibition are complicated and depend on both time and substrate. Satoh (14) has used the immobilized apoenzyme [Zn-free APase] for microanalysis of Zn(II) ions. This system could be used at least 120 times; treatment with a chelating agent regenerated the reactor following each measurement. Maniatis et al. (1) used EDTA together with high temperature to terminate the dephosphorylation reaction.
50 140 120 100 .,..4 9
p. 80
< 0
60
.,.,.1 l,.,,,d
40
o
GdCI
o
Urea
20 I
I--
1
2
I
I
I
3
4
5
C oncentration(M
)
Fig. 1. Effects of denaturant on APase activity.
100
p.
80
< 0
r .!.=,
60
40
: 20
"--.... 0
10
20
30
EDTA(~tM)
Fig. 2. Effects ofEDTA on APasr activity.
40
50
60
70
51 120 100
80
~-
e A
0
<
t~
".=.
1,10 Phenanthroline 4,7 P h e n a n t h r o l i n e
60 40
0
5
10
15
C oncentration(m
20
M )
Fig. 3. Effects of Phenantrolines on APase activity.
120 100 -~9
80
<
60
A m
,ab
o.,,t
,..,
o
40
THF
m
20
DMF
A
0
20
DMSO
40
%
A 60
v/v
Fig. 4. Effects of Solvents on APase activity.
solvent
80
100
52
Effect of Solvents. APase showed great tolerance of the solvents DMF, DMSO and THF as their concentrations (% v/v) increased (Fig. 4). APase completely withstands up to 60% (v/v) DMSO (with a slight activation effect); above 63%, activity declines in a threshold manner and is absent at 66%. Similarly, APase is completely tolerant of DMF up to 66% (v/v). Here also, a steep threshold effect occurs and activity is virtually absent at 80% DMF. THF has no effect on activity up to 82% (v/v) and approximately 30% activity is retained in 100% THF. This last result is remarkable in view of THF's high denaturation capacity value for enzymes (15). Effect of SDS. Low SDS concentrations adversely affect APase; however, more than 50% of initial activity remains at 1% (w/v) SDS and approximately 30% activity is retained at 10% (w/v). Clearly, APase retains significant activity at the 0.5% level of SDS used by Nakagami et al. (10) in their prehybridization mixture. This study helps to define tolerable storage and reaction conditions for native APase; use beyond these limits will likely require the implementation of one or more stabilization strategies (reviewed in 16). Boivin et al. (17) have described the stabilization of calf intestinal APase by immobilization and chemical modification. Acknowledgement. We thank Mr Damien O'Brien for help in preparing this manuscript. REFERENCES 1. Maniatis T, Fritsch EF, Sambrook J. Molecular Cloning: a Laboratory Manual. Cold Spring Harbour, NY: Cold Spring Harbour Laboratory, 1982: 133-134. 2. Harris MR. Biotechnol Adv 1991; 9:185-196. 3. Bronstein I, Voyta JC, Thorpe GH et al. Clin Chem 1989; 35:144 l- 1446. 4. Gray CH. Clinical Chemical Pathology, 7th edn. London: Edward Arnold, 1974:110. 5. Femley HN. In: Boyer PD, ed. The Enzymes vol 4. NY: Academic Press, 1971: 417-447. 6. Tietz NW, Rinker AD, Shaw LM. J Clin Chem Clin Biochem 1983; 21:731-748. 7. Pace CN, Shirley BA, Thompson A. In: Creighton TE, ed. Protein Structure: a Practical Approach. Oxford: IRL Press, 1989:311-329. 8. Franks F. Trends Biotechnol 1994; 12" 114-117. 9. Yang B, Viscidi R, Yolken R. Anal Bioehem 1993; 213" 422-425. 10. Nakagami S, Matsunaga H, Oka Net al. Anal Bioehem 1991; 198" 75-79. 11. de la Foumiere L, Nosjean O, Buchet R et al. Bioehim Biophys Acta 1995; 1248: 186192. 12. Chakrabarti A, Basak S. Eur J Biochem 1996; 241" 462-467. 13. Czekay C, Bauer K. Biochem J 1993; 290: 921-926. 14. Satoh I. Biosensors Bioelectron 1991; 6: 375-379. 15. Khmelnitsky YuL, Mozhaev VV, Belova ABet al. Eur J Biochem 1991; 198: 31-41. 16. O F~gain C. Stabilizing Protein Function. Berlin: Springer Verlag, 1997. 17. Boivin P, Kobos RK, Papa SL et al. Biotechnol Appl Biochem 1991; 14: 155-169.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
53
Stabilization o f ~-glucanase and its effect on the substrate h y d r o l y s i s pattern I. Markovi6, B. Markovi6-Dev~i6, S. Gamulin and N. Pavlovi6 PLIVA d.d., RESEARCH INSTITUTE, Prilaz baruna Filipovi6a 25, 10000 Zagreb, Croatia
The enzyme preparations for barley 1,3;1,4-13-D-glucan degradation are complexes of several 13-glucanases with endo- and exo-type activities. During stabilization procedure, the enzymes are subjected to environmental changes such as temperature, ion strength etc. They could alter the proportion of enzymes, and consequently, produce certain changes in substrate hydrolysis pattern. Thus, the effect of NaCI and of spray-drying on [3-glucanase (Aspergillus niger) storage stability as their influence on the pattern of the substrate hydrolysis was studied. Possible implications of changed hydrolysis pattern, caused by stabilization procedure, on the efficiency of the enzymes used for specific purposes were discussed.
1. INTRODUCTION Various enzymes are involved in degradation of barley 1,3;1,4-13-D-glucan ([3-glucan): endo-l,4-13-glucanase (EC 3.2.1.4), exo-l,4-~-glucanase (EC 3.2.1.91), endo-l,3;1,4-[3glucanohydrolase lichenase (EC 3.2.1.73), 13-glucosidase (EC 3.2.1.21) and possibly some other enzymes [ 1.2]. Enzymatic degradation of [3-glucan is very important because of its effect on the industrial exploitatior: of bad.:y g~"Zn. Enzyme preparations from various sources show different patterns of I~-giucan hydrolysis: ,:3me enzyme complexes exhibit endo-type action and others act more exogenously [3,4]. Generally, the type of action depends on the proportion of each enzyme in an enzyme complex. The studies of physico-biochemical changes of [3-glucanase during storage has shown that 13-glucanase activity decreases whereas 13-glucosidase activity is not significantly changed [5]. Similar results have been obtained in cellulase storage [6]. In the methods used for enzyme stabilization, active molecules are often subjected to marked environmental changes (increased ion strength, thermal stress etc.). These can change biochemical characteristics of enzyme complex and thus favourably or unfavourably affect the enzyme properties. Taking this into account, we studied the effect of NaC1 concentration and spray-drying on 13-glucanase storage stability. The influence of these methods for enzyme stabilization on the substrate hydrolysis pattern was also examined.
54 2. MATERIALS AND METHODS The enzyme complex for barley 13-glucan degradation was produced by submerged cultivation of Aspergillus niger [7]. The culture filtrate was concentrated on the rotary vacuum evaporator, Jedinstvo, Zagreb, Croatia (at 45 ~ and 2.7 kPa) to obtain 7% of dry matter. Appropriate quantities of NaCI was added for stabilization of enzymatic activity to the concentrated enzyme preparation (10 units/mL; pH 4.85). To assess enzyme stability, the enzyme preparations were kept in brown reagent bottles at 4 ~ and at 25 ~ (room temperature) respectively. To obtain powdered ~-glucanase, the culture filtrate was evaporated to 18-20% dry matter; NaCI 15% and sorbitol 8% were added for stabilization. Spray-drying was optimized in a Btichi 190 Mini Spray Dryer, Flawil, Switzerland. A/S NIRO ATOMIZF.R, Copenhagen, Denmark pilot-plant spray-dryer was used for enzyme production. Inlet and outlet air temperature was 185 ~ and 85 ~ respectively and delivery rate of enzyme solution 9-11 L/h. One unit of [3-glucanase activity is the amount of enzyme which liberates reducing carbohydrates with reduction power corresponding to 1 grnol glucose per minute from 0.5 % solution of I]-glucan at 30 ~ and pH 5.0 (1/30 M phosphate buffer [4]. One unit of 13-glucosidase is defined as the amount of enzyme which liberates 1 ~tmol p-nitrophenol from p-nitrophenyl-13-D-glucopyranozide within 10 minutes at pH 4.5 (0.05 M acetate buffer) and 45 ~ [8]. For the study of [3-glucan hydrolysis pattern, the substrate was prepared by cooking in water over 10 minutes. Solubilized 13-$1ucan (1% (w/v)) was incubated with 0.03 units/mL in a water bath shaker (mixing 100 min l ) at 50 ~ and pH 5.0 (0.1 M acetate buffer). In the 13-glucanase activity determination and studies of 13-glucan degradation kinetics, reducing sugars (as glucose) were determined according to the Somogyi-Nelson method [9]. Barley I]-glucan was purchased from Sigma, St. Louis, U.S.A. Total carbohydrates were determined as described in the literature [10]. Conversion was defined as ratio reducing sugars/total carbohydrates multiplied by 100. Glucose was determined by the GOD-PAP method on an automatic analyser (TRAACS 800, BRAN+LUEBE, Norderstedt, Germany). Cellobiose and higher oligosaccharides were analysed by thin layer chromatography [4].
3. RESULTS AND DISCUSSION 3.1. Stabilization with NaC! Neutral salts stabilize enzymes so that their ions compete for water or bind charged groups. This results in an increased enzyme molecule resistance to unfolding reactions. At high concentrations due to their osmotic effect, salts prevent microbial growth and digestion of enzyme molecules [ 11 - 13]. The effect of NaCI, in the concentration range of 5-20%, on the stability of liquid ~-glucanase at 4 ~ and 25 "C was examined. Although enzyme deactivation is a very complex process, it could be analysed by means of first-order kinetics [14]. Added NaCI stabilized liquid [3-glucanase preparation in long-term storage. The enzyme storaged at 4 ~ retained its activity for 12 month regardless of NaCI concentration used. At 25 *C the enzyme decay constant decreased from 1.6 x 104 h -t (control) to 4.7 x 10.6 h -I for the enzyme preparation stabilized with 20% NaCI (Fig. 1). Enzyme decay exhibited biphasic character. Thus, in NaC1 concentration range of 5-10% the enzyme decay function followed the firstorder kinetics during 1920 h (3 month) storage. Under the prolonged storage, the enzyme
55 decay constants increased. Similar results were obtained for enzyme stabilization with 12.5% and 15.0% NaC1. Deviation from the first-order kinetics, however, occurred after longer storage (5760 h or 8 months). That type of deactivation was in accordance with the proposed model that assumed microbial contamination [14]. In the NaCI concentrations of 17.5% and 20.0%, the enzyme retained practically entire activity during 12 month storage. It was not recorded that stabilization with NaCI had significant effect on the substrate hydrolysis pattern.
0.5 0.0
4
-0.5 ~" ,,-n -1,0 v
c
-1.5 -2.0 -
O
-2.5 -
Control
9
5% NaCI
-3.0 0.5
0.0 -~....__,-,__
~
. . . . . . . . . . . . . . . . . . .
-0.5 O UJ
-1.0 -
,,'1 -1.5 -
O
i--
-
-2.0
-
-2.5
-
7.5% NaCI 9
v
10% NaCI 12.5% NaCI
-3.0 0.5
0.0 0 U.l
i~
-(
"
"
"
L:..:_L
. . . . .
L L L L "
"
-0.5 -1.0
-
-1.5
-
-2.0
-
vr
--
0
-2.5 -3.0 0
15% NaCI 9
17.5% NaCI
v
2 0 % NaCI I
I
I
I
I
I
I
I
1
2
3
4
5
6
7
8
9
"Rme (h) x 10 "~
Figure 1. Plot of In E/E0, t for deactivation of I~-glucanase depending on NaC1 concentration. E0, E - Enzyme activity at beginning of storage (10 units/mL) and after time t. Temperature: 25 ~ pH 4.85.
56
3.2. Spray-drying Enzyme solutions are often dried to produce stable enzyme or to prepare for application convenient enzyme form. Spray-drying offers economical advantages for the production of enzyme in a powder form [15]. However, during spray-drying, enzyme molecules are subjected to thermal stress followed by an enzyme deactivation. To reduce the loss in enzyme activity, various additives are used (sugars, polyols, salts). In order to prepare a spray-dried ~-glucanase, the addition of NaCl and sorbitol was optimized. Taking into account losses in enzyme activity and the capacity of the sprayer, the addition of 15% NaCl and 8% sorbitol was found to be the most acceptable. Overall ~-glucanase activity losses during drying were in the range of 25-40%. Dried enzyme showed high storage stability; after 12 month storage the enzyme activity was not significantly reduced. However, due to thermal stress, the ratio of enzymes in the enzyme complex appeared to be changed. That was indicated by the decrease of ~-glucosidase activity and changes of [3-glucan degradation kinetics. Thus, specific [3-glucosidase activity during spraydrying decreased from 136.2 units/mg protein in liquid 13-glucanase to 20.8 units/mg protein in spray-dried ~-glucanase. In addition, in ~-glucan hydrolysis with liquid and spray-dried ]3-glucanase (0.03 units/mL) the conversions during initial 20 rain were very close. After that time, however, ]]-glucan hydrolysis with spray-dried 13-glucanase proceeded faster than in liquid.13-glucanase. Conversion degree of [3-glucan degradation with spray-dried [3-glucanase after 120 min hydrolysis was 20% and 15% in hydrolysis with liquid ~-glucanase preparation (Fig. 2). 25
9Spray-dried ~glucanase o Liquid I~-glucanase 20
~c -1 5 O W 0
"10
0
tO
i~
!
!
!
!
!
i
!
0
20
40
60
80
100
120
140
160
Time (rain) Figure2. Kinetics of I]-glucan hydrolysis with liquid and spray-dried 13-glucanase preparation. Substrate concentration: 1% (w/v); enzyme concentration: 0.03 units/mL; temperature 50 ~ pH (0.1 M acetate buffer): 5.0
57 Accordingly, glucose, cellobiose and other oligosaccharides concentrations were higher during hydrolysis with the spray-dried ~l-glucanase than in hydrolysis with liquid ~l-glucanase (Table 1). Table 1. Kinetics of products liberation during ~-glucan hydrolysis with liquid and spray-dried [3-glucanase preparation. (Conditions: see Fig. 2) Time (min) Degradation products (mg/mL) ~l-Glucanasepreparation 20
40
60
80
100
120
140
160
Liquid
0.002 0.004 0.008 0.009 0.014 0.016 0.019 0.023
Spray-dried
0.004 0.006 0.009 0.015 0.016 0.022 0.024 0.028
Liquid
Trace Trace Trace 0.107 0.133 0.16 0.16 0.187
Spray-dried
Trace 0.16 0.187 0.16 0.213 0.267 0.347 0.4
Liquid
0,133 0,24 0.347 0.373 0.56 0.747 1.173 1.28
Glucose
Cellobiose
*DP 3 Spray-dried
0,213 0,32 0.613 0.827 1.04 1.307 1.573 1.627
Liquid
Trace Trace Trace 0.08 0.107 0.16 0.16 0.187
Spray-dried
0.107 0.133 0.133 0.213 0.24 0.347 0.4 0.427
*DP 4 *DP- degree of polymerization In other words, spray-dried [3-glucanase showed more expressed exo-action pattern than liquid ~-glucanase. Analysis of kinetic data also proved that during spray-drying 13-glucanase catalytic characteristics altered. [3-Glucan hydrolysis using liquid [3-glucanase followed firstorder law, while kinetic data for ~-glucan hydrolysis using spray-dried I]-glucanase showed linear relationship in Foster-Niemann plot [ 16]. That suggested that the process inhibition by the reaction products was more expressed during hydrolysis with spry-dried [3-glucanase than in the case of liquid [3-glucanase. The alterations of catalytic characteristics during spray-drying of enzyme complexes should be considered in the light of their effect on the results of enzyme use. For instance, [3-glucanases in brewing are used to improve beer filtration. One may expect that they would not interfere with qualitative properties of beer. Accordingly, the release of higher quantities of fermentable glucose in spray-dried ~-glucanase may be an undesired feature of enzyme, from the point of its application in brewing. This may be particularly important in the production of alcohol-free beer [4]. The results of enzyme application in animal feeds are likely to be the most susceptible to mode of enzyme action. According to recent studies on physiological mode of feed enzymes action, the effect of "digestion kinetics" has major impact on the overall effects observed [17]. Thus, in the selection of the procedures for stabilization of enzyme complexes, besides on overall activity yield, one should take into account possible alteration of catalytic characteristics and their effect on the efficiency of the enzymes used for specific purposes.
58 4. CONCLUSION In selection of the procedures for stabilization of enzyme complexes involved in biopolymer degradation, it is necessary to take into account possible alteration of catalytic characteristics and their effect on the efficiency of the enzymes used for specific purposes. REFERENCES
1. B.V. McClear and M. Glennie-Holmes, J. Inst. Brew., 91 (1985) 285. 2. T. Kanda, H. Yatomy, S. Makishima, Y. Amano and K. Nisizawa, J. Biochem., 105 (1989) 127. 3. D.J. Manners and G. Wilson, Carb~.hydr. Res., 37 (1974) 9. 4. I. Markovi6, B. Markovi6-Dev~.i6 and 3. Joveva, Monatsschr. Brauwiss., 44 (1991) 312. 5. K.G. Gali6, I. Markovi6, B. Markovi6-Dev~i6 and N. Cikovi6, Monatsschr. Brauwiss., 42 (1989) 113. 6. G. Schaffeld, A. Illanes and L. Nunez, MIRCEN-J. Appl. Microbiol. Biotechnol., 4 (1988)414. 7. B. Markovi~-Dev~i6 and I. Markovi~, Postupak za pripravu enzima 13-glukanaze, Yugoslav Patent No 44537 (1985). 8. R.F.H. Dekker, J. Gen. Microbiol., 127 (1981) 177. 9. N. Nelson, J. Biol. Chem., 153 (1944) 375. 10. I. Markovi6, R. Deponte, V. Mari6 and V. Johanides. Process Biochem., 30 (1995) 411. 11. ~. Jane~ek, Process Biochem., 28 (1993) 435. 12. L. Gianfreda and M. R. Scarfi, Mol. Cell. Biochem., 100 (1991) 97. 13. C. O'Ffigftin, H. Sheehan, R. O'Kennedy and C. Kilty, Process Biocherrh, 23 (1988) 166. 14. J.P. Henley and A. Sadana, Biotechnol. Bioeng., 28 (1986) 1277. 15. N. Papamichael and H. Hustedt, Dechema Biotechnology Conferences 3, VCH Vedagsgesellschaft, Berlin, 1989. p 1127. 16. R.J. Foster and C. Niemann, Proc. Nat. Acad. Sci. USA, 39 (1953) 999. 17. A. J. Mul and A. W. Bonte, Proceedings of the 2nd European Symposium on Feed Enzymes (Eds: W. van Hartingsveldt, M. Hessing, J. P. van der Lugt, W. A. C. Somers), TNO Nutrition and Food Institute, Zeist, 1995, p 39.
Stability and Stabilizationof Biocatalysts A. Ballesteros,F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998ElsevierScience B.V. All rightsreserved.
59
S o m e factors affecting the behavior of anhydrous c~-chymotrypsin at high temperature Domenico Pirozzi Guido Greco jr.
Dipartimento di Ingegneria Chimica, Universit?l degli Studi di Napoli "Federico H", Napoli, Italia The influence of dehydration procedure and of KC1 addition on thermal stability of anhydrous (x-chymotrypsin was analysed. At high temperature (100 ~ formation of insoluble and inactive protein aggregates was observed. The partially inactivated proteins were characterised in terms of fraction of soluble enzyme and its specific activity. Secondary structure alterations of thermally treated, lyophilised proteins were characterised by FTIR spectroscopy. 1. INTRODUCTION Recently, significant progress has been achieved in understanding enzyme behavior in non-aqueous environments. The role of several key parameters, such as water thermodynamic activity [1], dehydration procedure [2] and colyophilised additives [3-4] on the structure and on the enzymatic activity and selectivity of anhydrous proteins has been analysed, as well. Little attention has been paid to the effect of such parameters on the stability of non-aqueous enzymatic systems at high temperature. The aim of this paper is to highlight some aspects of thermal inactivation phenomena of anhydrous proteins, pre-equilibrated in humidity-controlled environments. 2. MATERIALS AND METHODS
2.1 Materials and activity measurement Bovine pancreas (x-chymotrypsin (EC 3.4.21.1, Boehringer Mannheim) was purchased as a lyophilised powder and used without further purification. All reagents were obtained from Sigma-Aldrich S.r.l., and were analytical grade. Organic solvents were used after shaking with 3 ~ molecular sieves. Hydrolysis of 5 m g / m l N-succinyl-L-phenylalanine-p-nitroanilide (SUPHEPA) in 100 mM triethanolamine-HC1-NaOH buffer, pH 7.8 was carried out as the reference reaction. P-nitroaniline released during the enzymatic hydrolysis was monitored in a spectrophotometer, by direct reading at 405 nm.
60 2.2 Deactivation runs
12.5 m g / m l of (z-chymotrypsin solutions in 10 mM Na-citrate buffer, pH 3.00, were dehydrated following three different procedures. In the first, protein was lyophilised by freezing in liquid nitrogen and subsequent drying under vacuum for 72 h. In the second, c~-chymotrypsin was desiccated under vacuum in the presence of silica gel. The third procedure consisted in protein precipitation by PEG addition. The polymer was added until 40% (w/w), keeping the solution at 2~ under gentle stirring. Then, the solution was stirred 30 min, and proteins were recovered by centrifugation. Enzyme samples obtained by each dehydration procedure were inserted in test-tubes and kept for at least 7 days under vacuum, over silica gel, in order to achieve a constant thermodynamic activity of the hydration water. At the end of the equilibration process, the test-tubes were sealed and placed into a thermostated bath. Samples were drawn at predetermined time-intervals. Thermogravimetric analyses were carried out (Du Pont 951 TGA) on solidstate enzyme samples prepared according to the same general procedures described above. 2.3 S o l u b i l i t y determinations
After the exposure at high temperature and subsequent cooling to r o o m temperature, the enzyme was redissolved by injection of 1 ml of 100 m M triethanolamineHC1-NaOH buffer, pH 7.8 (optimal pH for enzyme activity) into the test tube, followed by gentle stirring for 70 min. The undissolved protein was then removed by centrifugation. Residual protein concentration in aqueous solution was determined by the BCA method. The extinction coefficient was found to be insensitive to modifications undergone by the enzyme. 2.4 FTIR tests
FTIR spectra of solid-state 0~-chymotrypsin after different exposure times at 100 ~ were measured by using a Mattson 5020 system, equipped with a DTGS KBr detector. For each sample, a 50 scans spectrum was collected at a resolution of 2 cm q. Additive-containing samples were corrected for the background. Each sample was prepared by mixing 1 mg of protein powder to 100 mg of KBr. After homogenization with agate mortar and pestle, the powders were pressed into pellets by using a hydraulic press. Spectral data elaboration was carried out by MicroCal Origin 2.94 software. The number of components and their wavenumbers were determined by second derivative spectra, calculated with an l 1-point smoothing function. Results were used as starting parameters in Gaussian curve fitting, performed on original spectra to determine wavenumbers and areas of the component peaks. A linear baseline was selected. Obviously, regressions were performed without fixing peak wavenumbers, height and with at half-maximum of individual bands [5]. The following assignment of component peaks to secondary structural elements was made [6]: 0~-helices, 1654 cm-1; [3-sheets, 1627, 1637 and 1674 cm -1.
61 3. RESULTS AND DISCUSSION
3.1 Effect of dehydration procedure Thermally treated (100 ~ anhydrous proteins underwent insolubilisation phenomena. For any dehydration procedure, the insoluble fraction increased with increasing thermal treatment time. Catalytic activity of insoluble proteins was negligible, as compared to that pertaining to the soluble fraction. Consequently, we characterised protein denaturation phenomena both in terms of soluble enzyme fraction and of its specific activity. The effect of dehydration procedure on the t i m e c o u r s e of thermallyinduced insolubilisation at 100 ~ is reported in Fig. 1 (curves a, b, c), with reference to lyophilisation, vacuum-desiccation and PEG-precipitation, respectively. Obviously, all experimental curves start from 100%, since no insolubilisation occurs in the absence of thermal treatment. After 100 hours, over 50% of proteins in each enzyme sample undergoes insoluble aggregation. Lyophilized proteins show faster aggregation kinetics than vacuum-desiccated and PEG-precipitated samples. I
s
o
0,8
!
i
i
!
........................
{ ..................................
-- .....................
i
i
i
i
!
I
!
i
~ .................................
!
i
i
{ ...................
r
0,6 0
0
0,4 (o(d) .......
0,2 i ..............................i.........................
~
k
0
,
i
i
,
!
50
i
I
(
I
a I
100
time, h
I
) I
!
~ I
150
(b) J
i
I
f
200
Figure 1. Insolubilisation curves of solid-state 0~-chymotrypsin; lyophilised (@), vacuum-desiccated (A), PEG-precipitated ( , ) e n z y m e ; 40% KCl-containing (w/w) lyophilised enzyme (O). Enzymatic activity of the soluble o~-chymotrypsin fraction was measured in aqueous solution, in order to minimise the effects related to the potentially different amounts of residual water. Indeed, significant differences in terms of residual enzyme activity have been observed [7], depending on the activity tests having been performed in organic solvents or in aqueous solution. Fig. 2 (curves a, b, c) shows the deactivation profiles at 100 ~ All curves start from virtually the same activity value. This indicates that protein activity is
62 not affected in the course of dehydration, whatever the technique adopted. Samples obtained by PEG precipitation display a very poor stability, as compared to the deactivation profiles pertaining to the other dehydration techniques. 10.4
..... t ...... L ..... ................................ ................................
a. .....
2. .....
~ ..... J., .... ~. ..... ~. ................................. ..~ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1 ......
! ............
L ...... | ...... . ................................. 9. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
L .....
.L .....
4" ..... L ..... 1 ...... ..- ................................ ... ................................
J ......
! .....
A O~llq
10.5
O
1 0 "6
-!!!!!!!!!!!!!!!!!!!!!!!!!!!~!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!i!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!i!!i!!!!!!!!!!!!!!!!!!!!!!!!!!!!! ___:!!!!!!!!!!!!!!}!!!!!!:::::::::i::(c):::::!!!!!?!?!!?!!!!!!!!i?!!!: ................
10"7 0
50
100 time, h
150
200
Figure 2. Deactivation curves of soluble (z-chymotrypsin fraction; lyophilised (@), vacuum-desiccated (A), PEG-precipitated (~) enzyme; lyophilised enzyme containing 40% (w/w) KC1 (O). The water-content of anhydrous proteins after the pre-equilibration procedure and prior to thermal treatment was estimated by thermogravimetric analysis. Results are reported in Table 1 (samples 1, 2, and 3). It should be noted that the water content is expressed in terms of mass of water per overall mass of protein and buffering salts. As regards the data obtained with KC1, to be discussed later, the same reference holds since the amount of water contained in KC1 crystals is negligible. Table 1 Water content of enzyme samples prior to thermal treatment sample 1 2 3 4
dehydration procedure lyophilisation vacuum-desiccation PEG-precipitation lyophilisation, KC1 40% (w/w)
water content (% w / w ) 1.67 1.72 2.44 1.68
,
The deactivation rates are clearly related to the initial water-content of enzyme samples: the higher the latter the faster the deactivation. On the contrary, solubility-time curves cannot be correlated to initial enzyme hydration levels.
63 3.2 FTIR analysis of thermodenaturation phenomena of lyophilised proteins Secondary structure alterations undergone by the lyophilised enzyme upon thermal treatment were studied by FTIR spectroscopy in the amide I region. These experimental tests were not performed on enzyme samples dehydrated by the other techniques since protein homogenization was virtually impossible, unless potentially disruptive mechanical procedures were adopted. Tab. 2 (samples 1, 2, and 3) shows the results of the quantitative analysis of the FTIR spectra as a function of thermal treatment time at 100 ~ Table 2 Secondary-structure alterations after thermal treatment at 100 ~
secondary structure, % sample
time, h
additive
(z-helix
[~-sheet
others
1 2 3 4 5 6
0 24 72 0 24 72
none none none KC140% (w/w) KC140% (w/w) KC140% (w/w)
11.0 9.7 7.8 10.9 10.0 7.0
46.4 46.9 41 42.1 42.3 40.3
42.6 43.4 51.2 47 47.7 52.7
,,,
The bands originating from ~-sheet structural elements display a slight rise after 24 h exposure at 100 ~ followed by a drop after 72 h. As regards (z-helices, a significant decrease is recorded as exposure time increases. The alterations recorded in the first 24 h of thermal treatment (increase in 13sheets, decrease in (z-helices) are qualitatively similar to those produced by the lyophilisation process, as reported by Klibanov and co-workers [5,8]. In the mentioned papers, the authors explained the increase in ~-sheet content as an effect of the formation of intermolecular ~-sheet structures due to dehydration. Similarly, we can assume that, due to the thermal treatment, new intermolecular ~-sheets are formed. This is in agreement with the time-progressive protein aggregation observed in Fig. 1. After 72 h exposure at 100 ~ both (z-helices and [3-sheets decrease. This is clearly due to thermally-induced protein disruption. Again, this agrees with the progressive loss of catalytic activity reported in Fig. 2.
3.3 Effect of KC1 addition Additives are often included in enzyme preparations to protect proteins during dehydration [3,4]. Also, they were found to be effective against protein selfinteraction mechanisms [9]. Therefore the analysis of 0~-chymotrypsin thermal stability performed on samples co-lyophilised with KC1 might be of some interest. Addition of the inorganic salt was effective in slowing down the aggregation kinetics, as reported in Fig. 1 (curve d). This is not the case as regards the catalytic
64 activity of the soluble fraction due to a slight deactivating effect of KC1 (Fig. 2, curve d). Salt addition did not affect the water content of pre-equilibrated, anhydrous proteins, as shown in Tab. 1 (sample 4). The effect of KC1 on secondary structure alterations of lyophilised proteins exposed at 100 ~ was also studied. Results of the quantitative analysis made by FTIR spectroscopy in the amide I region are reported in Table 2 (samples 4, 5, and 6). A clear reduction of initial amount of ~-sheets was recorded on KC1additivated samples, as compared to proteins lyophilised without additive. This could be the reason why the kinetics of interproteic interactions is slowed down by KC1 (Fig. 2). Alterations recorded during thermal treatment are qualitatively similar to those pertaining to samples without KC1. An increase in ~-sheets and a decrease in (z-helices occurred in the first 24 h, whereas a fast disappearance of both types of secondary structures was detected for longer exposure times. This indicates that, even in the presence of KCI, early formation of new intermolecular ~-sheets is followed by a gradual disruption of the secondary structure of the protein. 4. CONCLUSION Formation of insoluble and inactive proteic aggregates can significantly reduce the operating stability of anhydrous (z-chymotrypsin at high temperature. Kinetics of aggregation and inactivation depend on the dehydration procedure followed. Enzyme thermal deactivation profiles are increasingly steep, the higher the initial water content of the protein. Aggregation phenomena are in agreement with the formation of new intermolecular ~-sheets observed by FTIR spectroscopy. Co-lyophilisation with KC1 slows down aggregation kinetics, although the stability of the soluble protein fraction is slightly reduced. REFERENCES
1. P.J. Halling, Enz. Microb. Technol., 16 (1994) 178-206. 2. H. Noritomi, A. Almarsson, G.L. Barletta and A.M. Klibanov, Biotechnol. Bioeng., 51 (1996) 95-99. 3. K. Dabulis and A.M. Klibanov, Biotechnol. Bioeng., 41 (1993) 566-571. 4. Y.L. Khmelnitsky, S.H. Welch, D.S. Clark and J.S. Dordick, J. Am. Chem. Soc., 116 (1994) 2647-2648. 5. K. Griebenow and A.M. Klibanov, Proc. Natl. Acad. Sci. USA, 92 (1995) 1096910976. 6. D.M. Byler and H. Susi, Biopolymers, 25 (1986) 469-487. 7. B. Schulze and A.M. Klibanov, Biotechnol. Bioeng., 38 (1991) 1001-1008. 8. P. Mishra, K. Griebenow and A.M. Klibanov, Biotechnol. Bioeng., 52 (1996) 609614. 9. W.R. Liu, R. Langer and A.M. Klibanov, Biotechnol. Bioeng., 37 (1991) 177-184.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
65
Horseradish p e r o x i d a s e stability in the course o f phenols oxidation A.M.Egorov, Yu.L.Kapeluich, T.A.Pastuhova, M.Yu.Rubtsova, D.N.Cherepanov Dept. Chem. Enzymology, Chemistry Faculty, M.V.Lomonosov Moscow State University, 119899 Moscow, Russia 1. INTRODUCTION Plant and microbial peroxidases are of great interest for various application in industry and research due to their wide availability and high activity in the reactions of oxidation of different organic substrates. Phenol and its derivatives belong to one of the most interesting class of peroxidase substrates from the practical point of view. Different methods of analytical biochemistry widely employ the reactions of phenols oxidation catalyzed by peroxidases [ 1-5]. The oxidation of phenols is also of interest for the purpose of obtaining new food dyes and pharmaceutical products [6]. It was shown that the radical polymerization of phenols can be applied to obtain the phenolic resins in large scale, to remove the chemical pollution from industrial waste water in the course of coal conversion, pulp and paper manufacturing, wood preservation, metal casting [7-11 ]. Generally the reactions catalyzed by peroxidases have radical mechanism. They, as a rule, include two stages: enzymatic formation of products, which are free radicals, and subsequent interaction of these radicals with various molecules in the reaction media [12]. As the process is accomplished by peroxidase inactivation [13, 14], the poor stability of the enzyme in the course of phenols oxidation can limit the industrial application of peroxidase. The enhanced chemiluminescence (ECL) reaction, i.e. the co-oxidation of luminol and a substrate (called an enhancer) by hydrogen peroxide in the presence of horseradish peroxidase (HRP), is one of the examples of peroxidase-catalyzed reactions. HRP inactivation in the course of the ECL reaction restricts the sensitivity of a number of analytical methods based on the usage of HRP as a label [15]. 2. RESULTS AND DISCUSSION In our previous study we established that HRP inactivation during the ECL reaction is induced by intermediate of oxidation of substituted phenol employed as an enhancer [ 16]. We determined the extent to which the enzyme can be inactivated during phenols' oxidation when the enzyme was incubated with fixed amounts of hydrogen peroxide and substituted phenol. Small aliquots of the inactivation mixture were taken periodically from reaction media to measure peroxidase activity towards 2,2'-Azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) . Figure 1 presents the data on peroxidase inactivation during oxidation of different substituted phenols investigated as potential enhancers for the ECL reaction. *Contract grant sponsor: Amersham International plc. Contract grant sponsor: ISSEP
66
100
9
~
4-iodophenol 4-phenylphenol 4'-hydmxy-4-biphenyl-cadmxylic acid 4,4'-dihydroxybiphenyl
+
9e
40
9
-
,,, 9 "'
0
20
40
60
I'
80
'
100
I
120
"nn~n Fig.1. Time-dependence of HRP inactivation in the course of peroxidation of phenol derivatives. We revealed the increase of peroxidase inactivation rate when peroxidase was incubated with a mixture of 4-iodophenol and hydrogen peroxide in contrast with incubation of peroxidase with hydrogen peroxide by itself (Figure 2A). B
A
120 -
110 HRP+4-iodophenol
~-, 9 qp,,l
~
100.1
;.0 ........ o,;
9
80-
90-
~
80-
~ t~
60-
9
40
i ..... HRP+H202 ll..ll ....... ......... 9 ........ 9 . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9
40 -
~ ~
2 0 - ee. HRP+H202+4-iodop henol
g
0 0
9
9 odophenol
............................... , .............................. ~ ....
...
60 -
"~
100~ P ~
I
I'"
I
I
1
50
100
150
200
250
Time,min
30 20-
300
2000
4000
6000
8000
10000
[H202I/[HRPI
Fig.2. A HRP inactivation during the incubation of the enzyme with H202, 4-iodophenol, and a mixture of H202 and 4-iodophenol. B: Dependence of H R remaining activity on [H202]/[HRP] ratio.
But the amount of catalytic events which the enzyme can perform before its complete inactivation was higher when peroxidase was incubated with a mixture of substrates (Figure 2B). The number of catalytic events before complete enzyme inactivation was determined by
67 extrapolation to complete loss of HRP activity from data of Figure 2B.The enzyme can make up to 10 000 cycles in the presence of hydrogen peroxide and 4-iodophenol and only 600 cycles in the presence of hydrogen peroxide alone, setting the conversion of 3 molecules of hydrogen peroxide during one catalytic cycle in the course of incubation of enzyme with hydrogen peroxide alone [17]. This notes the differences in inactivation mechanisms for both cases.
The phenoxyl radicals can be considered as one of the main intermediates of the reaction of phenols oxidation by hydrogen peroxide [12]. The generation of reactive oxygen species during this reaction was also discussed [ 18-20]. One can suggest the aromatic amino acids as the primary target of the attack of active oxygen. The possible targets on a protein globule for active oxygen radicals are aromatic amino acids. In this case the excess of phenolic compounds in the reaction media might prevent the etm3nne from inactivation. Figure 3 shows the protective role of 4-iodophenol on the inactivation as a result of action of sulfate radicals forming at the photodestruction of sodium persulfate. HRP+Na2S208+4-iodop henol
100
I;
HRP+Na2S208
0
t
a ' ~
~ l'O Thrr, rntn
1'2 ' 1'4
1'6"
Fig.3. Effect of 4-iodophenol on HRP inactivation rate induced by sulfate radicals through Na2S208 photolysis. To elucidate the role of active oxygen species (hydroxyl radicals and superoxide anion) we studied the effect of radical scavengers on the rate of peroxidase inactivation in the course of 4-iodophenol peroxidation [16]. The addition of sodium formate and mannitol as OH scavengers to the ECL mixture did not reveal any effect on the rate of phenol oxidation or HRP activity decay rate [16]. Thus we can conclude that active oxygen radicals are not the main inactivating agents in the reaction of phenols oxidation. Phenoxyl radicals present the other type of intermediate. In our previous work we established that if they are inactivating agents, they have to leave the active site of peroxidase before the enzyme will be inactivated [ 16]. Figure 4 presents the effect of BSA on the HRP inactivation rate in the course of 4-iodophenol oxidation. The effect consists in the increase of enzyme stability up to 20 fold.
68 40 30 t,-'.
"~ 20 10
I
0,0
9
i
0,2
9
I
0,4
.
I
0,6
9
I
0,8
9
i
1,0
--.
[BSA], mg/ml Fig.4. The dependence of half-time of HRP inactivation during the oxidation of 4-iodophenol on BSA concentration. If we suggest the loss of enzyme activity owing to non-specific interaction of radical species with protein globule, the stabilization role of BSA consists in the reaction of these species with excess BSA and not with amino acids involved in the active site or peripheral part of peroxidase globule. To examine the possibility for phenoxyl radicals to bind with the protein globule of peroxidase we studied the interaction of peroxidase with 4-hydroxycinnamic acid labeled with biotin (pHCA-Bt). pHCA-Bt as substituted phenol can be oxidized by hydrogen peroxide in the presence of peroxidase, and phenoxyl radicals covalently bound to biotin are formed as intermediates during the reaction. If they can react with the peroxidase globule, the peroxidase labeled with biotin can be obtained as a result of the enzymatic reaction. The biotin residues can be revealed then with a conjugate of streptavidin-peroxidase. To separate the products of the reaction we immobilized the peroxidase in the wells of polystyrene microtiter plates (Fig 5). A
...............
pHCA,Bt
.,,,.m,
~,m..,m|..1.,.1.,u,.mm
]~
w.l.t
Bt
Fig. 5. Schemes for the reactions of immobilized peroxidase with the products of pHCA-Bt peroxidation. On the first stage (Figure 5, line A) the immobilized peroxidase catalyses the oxidation of pHCA-Bt with H202, then, after a washing step, immobilized peroxidase supposedly modified with products of pHCA-Bt oxidation interacts with conjugate streptavidinperoxidase. Lines B and C show the control experiments. Line B is the control without oxidation of pHCA-Bt. Line C is the control without interaction with streptavidin-peroxidase conjugate.
69 After the experiments were performed accordingly the schemes A, B, C we determined the overall peroxidase activity towards ABTS in different wells from kinetic data (Figure 6). The H R activity after the oxidation of pHCA-Bt (curve C) was lower by 20% when compared with the activity of immobilized HRP (curve B), while the overall HRP activity after oxidation of pHCA-Bt followed by the interaction with streptavidin-HRP conjugate was higher by a factor of 2 (curve A). When the interaction with conjugate pHCA-Bt was performed in the presence of excess BSA (5mg/ml), the overall HRP activity was comparable with the activity of immobilized HRP (data not shown).
C~4
A
(13 o
Q2-
B
C
0.1-
%
10
20
30
40
Tirn~ rrin
50
60
7o
Fig.6. Kinetic curves for ABTS oxidation by peroxidase modified with products of pHCA oxidation. Curves A, B, C correspond to the schemes presented in Fig.5. Thus, the radicals of pHCA-Bt can interact with amino acid residues on the surface of peroxidase globule, and in the presence of BSA they completely bind with soluble BSA. The binding of phenoxyl radicals with protein globule followed by the modification of certain amino acids present on the surface of the protein may be the reason for HRP inactivation. The formation of soluble complexes of peroxidase and polycations was found recently [21 ]. We established the decrease in peroxidase inactivation rate at the addition of increasing amounts of polycation 2,5-ionene bromide. The time for half-inactivation was 60-fold higher at polycation concentration of 1 mM [ 16]. As 2,5-ionene has not any group whichmight interact with phenoxyl radicals we can propose that polymer can cover the protein globule with protective coating, which prevents surface amino acids from the interaction with phenoxyl radicals. The ability of phenyl radicals forming at peroxidation of phenylhydrazyne to modify protein globule and also to modify heme at 5-meso position and to oxidize 8-CH3 group was shown by Ator and De Montellano [22]. The feature of these substrates in comparison with the substituted phenols studied in the present work is that the number of catalytic events is lower by a factor of 1000. Thus, the strategy for HRP protection from inactivation in the course of enzymatic oxidation of phenols should be directed to the enzyme protection from the action of phenoxyl radicals when is going to be used for analytical applications. This can be realized by
70 the enclosing of protein or by site-directed mutagenesis. For industrial application of the enzyme it is possible to use different additives which might interact with intermediates and products resulting in the neutralization of inactivating action [23]. REFERENCES
1 2 3 4
5 6 7 8 9 10 11 12 13 14 15
16 17 18 19 20 21 22 23
Thorpe, G.H.G. and Kricka, L.J. (1986) in Bioluminescence and Chemiluminescence. Part B. Methods in Enzymology, v. 133 (DeLuca, M.A. and McElroy, W.D., eds.), pp. 331-353, Academic Press, Inc., Orlando. Kricka, L.J. and Thorpe, G.H.G. (1986) Parasitol.Today 2, 123-124. Kovba, G.V., Rubtsova, M.Y. and Egorov, A.M. (1997) J. Bioluminesc. Chemiluminesc. 12, 33-36. Rubtsova, M.Y., Gavrilova, E.M. and Egorov, A.M. (1994) in Bioluminescence and Chemiluminescence (Campbell, A., Kricka, L. and Stanley, P.E., eds.), pp. 365-370, John Willey & Sons, Cambridge. Osipov, A.P., Zaitseva, N.V. and Egorov, A.M. (1996) Biosensors and Bioelectronics 11, 881-887. Kvaratskhelia, M., Winkel, C. and Thorneley, R.N.F. (1997) Plant Physiology 114, 1237-1245. Klibanov, A.M., Tu, T.-M. and Scott, K.P. (1983) Science 221,259-260. Ayyagari, M., Akkara, J.A. and Kaplan, D.L. (1996) Acta Polymerica 47, 193-203. Dec, J. and Bollag, J.M. (1990) Arch. Environm. Contamination and Toxicol. 19, 543550. Miland,E., Smyth, M.R. and Fagain, C.O. (1996) J. Chem. Technol. Biotechnol. 67, 227-236. Tatsumi, K., Wada, S. and Ichikawa, H. (1996) Biotechnol. and Bioeng. 51, 126-130. Yamazaki, I., Mason, H.S. and Piette, L. (1960) J. Biol. Chem. 235, 2444-2449. Buchanan, I.D. and Nicell, J.A. (1997) Biotechnol. and Bioeng. 54, 251-261. Nicell,J.A. (1994) J. Chem. Technol. Biotechnol. 60, 203-215. Egorov, A.M., Kim, B.B., Pisarev, V.V., Kapeluich, Y.L. and Gazarian, I.G. (1993) in Bioluminescence and Chemiluminescence (Szalay, A., Kricka, L. and Stanley, P., eds.), pp. 286-300, John Wiley & Sons, Banff. Kapeluich, Y.L., Rubtsova, M.Y. and Egorov, A.M. (1997) J. Bioluminesc. Chemiluminesc. 12, 299-308. Arnao, M.B., Acosta, M., Del Rio, J.A., Varon, R. and Garcia-Canovas, F. (1990) Bioch. Biophy. Acta 1041, 43-47. Ma, X. and Rokita, S.E. (1988) Biochem. Biophys. Res. Comm. 157, 160-165. Dordick, J.S., Klibanov, A.M. and Marietta, M.A. (1986) Biochemistry 25, 2946-2951. Ortiz de MonteUano, P.R. and Grab, L.A. (1987) Biochemistry 26, 5310-5314. Gorovits, E.L., Izumrudov, V.A., Pisarev, V.V., Gavrilova, E.M. and Egorov, A.M. (1995) Biotechnol. Appl. Biochem. 22, 249-260. Ator, M.A. and Ortiz De Montellano, P.R. (1987) J. Biol. Chem. 262, 1542-1551. Nakamoto, S. and Machida, N. (1992) Water Res. 26, 49-54.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
71
The effect o f impurities of crude olive residue oil on the operational stability o f the Candida rugosa lipase i m m o b i l i z e d in p o l y u r e t h a n e foams Correia, A.C.; Ferreira-Dias, S. Instituto Superior de Agronomia, Departamento de Agro-Indtistrias e Agronomia Tropical, Tapada da Ajuda, 1399 Lisboa-Codex, Portugal
1. INTRODUCTION The current process "Colgate-Emery", for hydrolysis of fats and oils, is carried out in the absence of catalysts, at high temperature (higher than 250~ and under high pressure (50atm). The potential for the application of lipases (acylglycerol acylhydrolases, EC 3.1.1.3.) in the oleochemical industry is enormous. In lipase-catalysed reactions, a reduction in energy consumption and an increase in product purity are observed, due to the high selectivity and ability of these biocatalysts to work at room temperature. However, the current price of lipases is about an order of magnitude higher than the necessary to match the energy costs associated with standard processes ~. An immobilized lipase preparation with a high operational stability will reduce costs, making lipasecatalysed reactions a feasible process option. In addition, the inhibitory effect of several compounds such as free fatty acids (FFA) 2, chlorophyls, polyvalent metal cations (Cu, Fe and Ni) and oxidation products 3'4'5'6, on lipase activity, has been a limiting factor to the use of crude oils in lipase-catalysed processes. Therefore, the use of refined, bleached and deodorized (RBD) oils usually increases lipase activity and stability. However, refining additional costs must be considered and compared to biocatalyst costs 5. Encouraging results have been reported either for free lipases in the presence of an oil previously oxidized at 2050C7, or for the immobilized Aspergillus oryzae lipase in the presence of an industrial effluent of crude rapeseed oil in solvent media s . The characteristics of immobilization matrices are important on the operational stability of lipases. Successful results have been achieved with hydrophilic 9'1~ and hydrophobic supports i'll, as well as amphiphilic gels 12, on the hydrolysis, esterification and interesterification of oils and fats. The aim of this study was to investigate the operational stability of the Candida rugosa lipase immobilized in a hydrophilic biocompatible polyurethane foam during the batch hydrolysis of crude olive residue oil in n-hexane. This oil is extracted from olive husks, by solvent (n-hexane), after olive oil physical extraction. The crude olive residue oil may be used, after refining, for human consumption. The effect of different
72 contents of free fatty acids, pigments and oxidative products, of 2 crude olive residue oils on lipase stability was evaluated, for different amounts of water added. In addition, the effect of the removal of several impurities by adsorption to diatomaceous earths, alumina or activated charcoal, was also tested. The stability profiles of the biocatalyst, in the presence of the original crude and treated oils, were compared. 2. MATERIALS AND METHODS
2.1. Materials Crude olive residue oils containing 11% and 25% of free fatty acids (FFA), respectively, were a girl from Empresa Fabril de Moura, Moura, Portugal; granular (PK1) activated charcoal was donated by NORITT; diatomaceous earths, TONSIL. The liophylized lipase from Candida rugosa (lipase AY) was a gift from Amano, U.K.. The lipase was immobilized in a hydrophilic polyurethane foam (Hypol FHP 2002), kindly donated by Hampshire Chemical GmbH, Germany. Serum bovin albumin was from SIGMA, USA. All the other chemicals were of analytical grade and obtained from various sources.
2.2. Methods Immobilization: Lipase immobilization in the hydrophilic polyurethane foam occurred simultaneously with the polymerization 9. Foams were prepared by mixing 0.6 g of Hypol prepolymer with 0.3 g of aqueous phosphate buffer solution (0.020M KHEPO4+0.027M Na2HPO4; pH 7.0) and the lipase powder11~ Afterwards, these foams (with an apparent volume of about 4.9 cm 3 and a porosity of about 0.70) were cut into small cuboids (-43.07 cm 3) and introduced in the reaction medium. Hydrolysis Reaction- The immobilized lipase was added to a biphasic system consisting of (i) 12 cm 3 of an organic solution of crude olive residue oil in n-hexane and (ii) an aqueous phosphate buffer phase (0.020M KH2PO4+0.027M Na2HPO4) at pH 7.0 (molar ratio Water/Triglycerides = 4 or 19). The crude oil concentration used was calculated in order to have 60% (w/v) of triglycerides (TG) in solution, taking into account the difference in FFA content of the two oils. When previously used in the hydrolysis of refmed olive oil in n-hexane, this TG concentration promoted the activation of this lipase preparation 14. The reaction was carried out at 30~ in closed conical flasks, in a reciprocal water bath at 230 rev/min. Samples were taken out during the time-course of the reaction for the analysis of FFA. Initial rates were calculated by linear regression on the first 5 data-points (time, FFA concentration). Optimization of iipase loading- Different amounts of lipase powder ([mass of lipase/mass of pre-polymer] from 0.3 to 0.75) were immobilized inside polyurethane foams, prepared with 0.6g of pre-polymer, and tested during the hydolysis reaction (crude oil with 11% FFAcontent; water/TG=4). Batch Operational Stability Tests: The operational stability of the immobilized enzyme was assayed under similar conditions as previously described (c.f "Hydrolysis reaction"), with 350rag of lipase corresponding to a ratio lipase/pre-polymer (w/w) equal to 0.58. After 23 hour reaction time, the foams were removed from the reaction medium and rinsed twice with n-hexane (2 x 50 era3). One hour later (length of time required for the solvent to evaporate) the foams were introduced in fresh medium. The same lipase preparation was
73 reused for 10 consecutive days. Residual activities were daily evaluated by the assay of initial rates of hydrolysis. To investigate the effect of different amounts of water added, on lipase stability, experiments 1 and 2 were carried out with crude olive residue oil containing 25% FFA, and a ratio water/TG equal to 19 and 4, respectively. The effect on lipase stability of the removal of some inhibitory compounds, by adsorption to different adsorbents, was investigated. For experiments 3 and 4, the crude oil with 25% FFA content was solubilized in n-hexane (30%, w/v); a mass of diatomaceous earths, at a ratio earths/oil = 0.33, was added and allowed to contact for 24 hours, in a reciprocal water bath at 230 rev/min. The solvent was evaporated and one half of this oil was used in experiment 3. The remaining oil was treated with alumina (alumina/oil= 0.13, w/w) following the same protocol as with the earths. This final oil was used in experiment 4. For experiment 5, crude olive residue oil with 11% FFA content was used. For experiment 6, a solution of this original crude olive residue oil in n-hexane (30%, w/v) was previously passed through a chromatographic column filled with 30g of activated granular charcoal. The solvent was removed and the partially purified oil used. Analytical methods:. The FFA were assayed using the Lowry and Tinsley's colorimetric method is with benzene replaced by n-heptane 16. The content of chlorophyls was assayed spectrophotometrically by the absorbance of the solution oil in n-hexane at 668nm; oxidation products were assayed by u.v. spectrophotometry, at 232nm (hydroperoxides) and 270nm (final oxidation products, such as aldehydes and ketones). To determine if lipase loss occurred during batch reutilization process, protein content of the immobilized preparations was daily evaluated. Therefore, in parallel with batch operational stability experiments, 10 similar experiments were carried out (1 experiment corresponding to 1 different time). Prior to protein assay, the immobilized preparation was dried under vacuum, at 450C, for 8 hours. Afterwards, lml of NaOH 1N was added and placed in a water bath at 100~ for 5 min. Lowry et al.'s method 17 was used to evaluate protein content of this solution. Serum bovin albumin immobilized in the same support was used as a standard to establish the calibration curve. After treatment with granular charcoal, iron and copper in the original crude oil with 11% of FFA were assayed by graphite furnace atomic absorption spectrophotometry 18.
3. RESULTS AND DISCUSSION
Characterization of the oils- The oils were characterized in terms of oxidative products, pigments and Fe and Cu content (Table 1.). The use of adsorbents were effective on pigments removal. In contrast, a negligible adsorption of oxidative products was detected (Table 1.). A high reduction in copper content was observed on activated charcoal occurred, while iron content did not change (Table 1.). Optimization of lipase loading- A linear increase of the initial hydrolysis rate with lipase load was observed up to a weight ratio lipase/pre-polymer of about 0.6 (Fig.l). From this maximum on, reaction rate dropped as loading was increased. This indicates diffusioncontrolled limitations. In addition, no significant amounts of enzyme leakage occurred since changes in protein content were negligible (Fig.2). This was probably due to covalent binding between the active radical of the pre-polymer and amine groups of the enzyme, occurring simultaneously with entrappment during foam formation19
74 Table 1.- Characterization of the oils used in the experimenst- Absorbance values related to the presence of oxidative products (232 and 27Onto) and chlorophyls (668nm); Fe and Cu contents. (n.d.- not determined) . Experiment Oil Oil Abs Abs Abs Fe Cu Number FFA Treatment 232nm 270nm 668nm (ppm) (ppm) (%) ., 1, 2 25 11.453 6.057 4.940 n.d. n.d. 3
25
Earths
10.600
6.010
1.627
n.d.
n.d.
4
25
10.650
5.977
1.390
n.d.
n.d.
5
11
Earths+ Alumina -
11.807
10.963
6.893
18.11
4.20
6
11
Charcoal
11.757
10.857
0.197
18.11
0.19
50 (I)
t-
45 -
7,0
~
E-~ 4 0 -
"$
6,0
~e " L L 35
,', ~
5,0
ao-
~'~ 4,0
2225-g_.o_ "r" E 2 0 _
_ ~ 2,0
15
I
I
I
I
I
0,2 0,3 0,4 0,5 0,6 0,7 0,8 mass of lipase/mass of pre-polymer
Fig.l- The effect of lipase loading of polyurethane foams on the hydolysis rate of crude olive residue oil.
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i
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4"
i
i
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I
I
I .... !
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3
4
5
6
7
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Time (days)
Fig.2- The protein content of the immobilized lipase preparation during
I0 successive hydrolysis batches.
Batch operational stability- A fast deactivation of the bioeatalyst was observed for every system under study (Fig.3A). This disagrees with previous results where no deactivation was observed when the same lipase preparation was reused, in organic media, for 20 and 15 days, respectively, as a catalyst for the esterifieation of butyric acid with ethanol 9 and the glyeerolysis of olive oil l~ Concerning the present results (Fig.3- exp. 1 and 2), a higher stability was achieved under lower water content, suggesting that water may play a major role in the inactivation of the biocatalyst in the system under study.
75
B
A Relative activity
Application o f l u order inactivation model
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9
9
o 9
9
9
8
g
10
1
0
0
1
2
3
4
5
6
7
~rne (days)
Fig. 3- Batch operational stability tests to the polyurethane immobilized C. rugosa lipase during the hydrolysis of crude olive residue oil. A- Relative activity-data presented with reference to the initial hydrolysis rate of batch number 1 (1 batch = 23 hours). B- Application of l't order inactivation model. (Exp.1) 25% FFA crude oil, water/TG = 19.0; (Exp.2) 25% FFA crude oil, water/TG=4; (Exp.3) 25% FFA oil previously treated by activated earths, water/TG = 19.0; (Exp.4) similar to exp.3, but the oil was also treated by alumina; (Exp.5) 11% FFA crude oil, water/TG = 19.0;(Exp. 6) similar to exp.5, but the oil was treated by activated charcoal (see text for details).
76 When the 11% FFA crude oil was used (Fig.3- exp.5 and 6), the stability was lower than the observed with 25% FFA oil (Fig3-exp.l-4). This may be ascribed to the inhibitory effect of higher amounts of chlorophyls and secondary oxidative products in the first oil (Table 1). In fact, when pigments and copper were removed by adsorption to charcoal (exp.6), a slight increase in stability was observed as compared to experiment 5. Inactivation caused by FFA was not to be expected since the partition coefficient for oleie acid between the support and the organic solvent was close to zero 13. The removal of ehlorophyls from the more acidic oil with activated earths and alumina, also promoted an increase in lipase stability (Fig.3-exp.3 and 4). In the present study, the experimental data do not fit a first order exponential decay model (Fig.3B). The activity profiles suggest a non linear inactivation kinetics with sucessive inactivation stages 2~ for every experimental conditions tested. Further studies have to be carded out to understand the inactivation process caused by the impurities in crude oils.
Aknowledgements The authors are grateful to Prof. F/ttima Peres, Escola Sup. Agrhria de Castelo Branco, Portugal, for the Fe and Cu determinations. This study was supported by grant n ~ JNICT
PBIC/C/AGR/2316/95, Portugal. REFERENCES
I Brady, C.; Metcalfe, L.; Slaboszewski, D.; Frank, D (1988) Jr. Am. Oil Chem. Soc. 65 917921. 2 Dtinhaupt, A., Lang, S., Wagner, F. (1992), Biotechnol. Lett., 14 (10) 953-958. 3 Wisdom, R.A., Dunnill, P., Lilly, M.D. (1985), Enzyme Microb. Technol., 7 (11) 567-572. 4 Linfield, W. M. (1988), In: Proceedings World Conference on Biotechnology for the Fats and Oils Industry, (T. H. Applewhite, ed.), Am. Oil Chem. Soc., Champaign, USA, 131-133. 5 Posorske, L.H., LeFebvre, G.K., Miller, C.A., Hansen, T.T., Glenvig, B.L. (1988), J. Am.Oil Chem. Soc., 65, (6) 922-926. 6 Wang, Y., Gordon, M.H. (1991), J. Agric. Food Chem., 39, 1693-1695. 7 Legier, V., del Guist, C., Comeau, L. (1994), Rev. Fr. Corps Gras, 41 (3/4) 45-52. 8 Kuncov/t, G., Malrterovfi, Y., Loveckfi, P. (1994), Biotechnol. Techn., 8 (8) 535-540. 9 F. Dias, S., Vilas-Boas, L., Cabral, J.M.S., Fonseca, M.M.R. (1991), Biocatalysis, 5:21-34. l0 Ferreira-Dias, S. and da Fonseca, M.M.R. (1995) Bioprocess Eng., 13 (6) 311-315. 11 Hoq, M.M.; Yamane, T.; Shimizu, S.; Funada, T.; Ishida, S.(1984), J.Am. Oil Chem Soc., 61,776-781. 12 Kang, S. T.; Rhee, J. S.(1989), Biotechnol. Bioeng. 33: 1469-1476. 13 Ferreira-Dias, S., da Fonseca, M.M.R. (1995), Bioprocess Eng., 12 (5) 327-337. 14Ferreira-Dias, S., da Fonseea, M.M.R. (1995), Biocatalysis and Biotransformations, 13 (2) 99-110. 15 Lowry, R.R.; Tinsley, I.J. (1976) ,/. Am. Oil Chem. Soc. 53: 470-472. 16 Ferreira-Dias, S.; Fonseca, M.M.R. (1993) Biotechnol. Techn. 7: 447-452. 17 Lowry, O.H., Rosebrough, N.J., Farr, A.L., Randall, R.J. (1951), J.BioLChem., 193, 265 18 Peres, M.F.P. (1995),Sobre a oxidar de azeite virgem catalisada por metais, Tese Mestrado Cirncia e Tecnologia de Alimentos, UTL, Lisboa.. 19 Fassett, D.W. (1963)In: Industrial Hygiene and Toxicology, Interscience Publ., vol.2, pp: 2032-2033. 20 Henley, J.P., Sadana, A. (1986), Biotechnol. Bioeng.,28:1277-1285
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
77
A p p r o a c h e s for i m p r o v e d i d e n t i f i c a t i o n o f m e c h a n i s m s o f e n z y m e i n a c t i v a t i o n M. Polakovi~,, P. Vr~ibel* and V. B~le] Department of Chemical and Biochemical Engineering, Faculty of Chemical Technology, Slovak University of Technology, Radlinsk6ho 9, 812 37 Bratislava, Slovak Republic
This publication reviews the results of the authors' studies dealing with the identification of mechanisms of enzyme inactivation. It presents both a critical assessment of traditional approaches of evaluation of enzyme inactivation kinetics from uniresponse experiments as well as the procedures of better design and evaluation of experiments which improve the analysis of inactivation mechanism and its corresponding kinetics. These methods are based on the simultaneous evaluation of inactivation experiments carried out at several temperatures or pHs. The inactivation mechanism is thus evaluated from the data set containing larger amount of information compared to a single inactivation curve when the influence of the factor of variation is unequivocally defined, for example, by the Arrhenius equation in the case of temperature or protonation and deprotonation equilibrium constants in the case of pH. It is demonstrated furthermore that the complexity of the mechanism of the thermal inactivation of free enzyme can be enhanced by utilising the combination of selective immobilization and multitemperature evaluation when the immobilization prevents some reactions in the ordinary inactivation pathway to occur. All methods are illustrated on case studies using yeast invertase.
1. DRAWBACKS OF CONVENTIONAL EVALUATION [1] Enzyme inactivation is generally explained as a chemical process involving several phenomena like aggregation, dissociation into subunits, or denaturation (conformational changes), which occur simultaneously during the inactivation of a specific enzyme [2,3]. However, the principal information for elucidating the kinetics of enzyme inactivation is obtained through the measurement of enzyme activity, which is determined as the rate of enzyme reaction at specified conditions. Consequently, enzyme activity gives the best quantitative information on the phenomenon of activity loss, but it is difficult to relate this to the changes occurring in the enzyme structure. The total enzyme activity, A, is generally equal to the sum of the activities of active enzyme forms, A i . T h e activity, a, is usually given as a relative value of the initial activity, Ao,
' Present address: KluyverLaboratory for Biotechnology, Department of Biochemical Engineering, Delft University of Technology, Julianalaan 67, 2628 Delft, The Netherlands
78
E Ai
E ?'ici (1)
A
2A, o 2r,c,o i
i
where Yi are the molar activities and c i are the molar concentrations of enzyme forms. Equation (1) reflects the methodological problems encountered in the analysis of the kinetics of enzyme inactivation. As the inactivation is represented by a set of reactions among different enzyme forms, for the evaluation of kinetics it would be highly useful to know the content of individual enzyme forms. Further complications during the analysis of inactivation kinetics are that the molar activities of enzyme forms are generally unknown. Thus, it is difficult to decide from the inactivation curve alone whether a reaction is reversible or irreversible, whether parallel reactions are taking place on the same enzyme form or not, or whether intermediates are active or inactive. It has been demonstrated how simple kinetic equations (mainly first-order kinetics) can disguise a more complex kinetic behaviour [4-6]. Different inactivation mechanisms of equal complexity can however correspond to the same integral activity-time relationship and the number of equivalent mechanisms increases exponentially with the number of parameters in the kinetic equation [1]. Thus, neither isothermal inactivation data exhibiting non-first order kinetics do not provide sufficiently consistent information upon which a particular mechanism of inactivation could be declared. A careful statistical analysis of extensive set of inactivation data found in literature showed that any pattern of enzyme inactivation behaviour (biphasic, grace-period or activation-period) can be described by a relatively simple exponential function [1]. Such functions represent lumped kinetic equations and any simple mechanisms which could be identified from them are dismissed if the validity of the mechanisms is tested at several temperatures [ 1].
2. MULTITEMPERATURE EVALUATION [7] If the kinetic parameters are evaluated using a simultaneous fit of the inactivation data obtained at several temperatures, this provides an opportunity to improve the analysis of the mechanism and kinetics of inactivation. The improvement of the analysis consists in: 1. the inactivation mechanism should be validated at each temperature; 2. the kinetic equation may contain parameters which should not change with temperature; 3. the parameters dependent on temperature should provide a consistent temperature relationship, for example, the rate constants could be expected to obey the Arrhenius equation. Figure 1 presents the inactivation of yeast invertase at four temperatures from 40 to 60~ charaeterised by the so-called biphasic behaviour. In order to measure the benefits gained by the evaluation of inactivation kinetics from multitemperature data, isothermal data have been evaluated as well. The most distinctive difference between the quality of isothermal and multitemperature evaluation was found at the isozyme mechanism and is presented in Fig. 1. This mechanism was unsuitable for describing the inactivation of yeast invertase at all temperatures simultaneously whereas the corresponding isothermal model fitted the individual inactivation experiments very well.
79
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- - ~d.
-O
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r - 0.6 >,~ :L-'_ .> o
<
0.4
~',
0.2
iO 40"cl
Iv ~ ' c l _!v6. O~
k."
" ~ ~ . . ~ . ~ .~~ -
0.0
-L 0
i
i
10
20
=
30 Time
t
I
_i
40
50
60
(rain)
Figure 1. Thermal inactivation of yeast invertase at pH 4.6. Comparison of isothermal and multitemperature evaluation of inactivation experiments demonstrated on the isozyme mechanism. The calculated activity profiles are represented by solid (isothermal evaluation) and dashed lines (multitemperature evaluation). Reprinted from Enzyme Microb. Technol., Vol. 20, P. Vrfibel, M. Polakovi6, V. ~tefuca and V. B~ileg, Analysis of mechanism and kinetics of thermal inactivation of enzymes: Evaluation of multitemperature data applied to inactivation of yeast izivertase, 348-354, (1997), with permission from Elsevier Science. The method of multitemperature evaluation of inactivation data was tested using a number of inactivation mechanisms [7]. It was found that the lumped character of inactivation kinetics inherent to the isothermal activity-time relationship was significantly weakened. Two-reaction models, which all provided the same values of the sum of squares at the isothermal evaluation, could be comfortably distinguished here. Even the three-reaction models could be meaningfully discriminated. Combining the results of statistical evaluation (Fig. 2) and available literature data on structural changes of invertase during inactivation, it was concluded that the inactivation of invertase took place in at least three reaction steps which could be the dissociation of oligomeric enzyme into subunits, denaturation and association both resulting in the complete loss of activity.
3. CONTRIBUTION OF SELECTIVE IMMOBILIZATION [81 A combined effect of selected immobilization techniques and the multitemperature evaluation of the inactivation of immobilized preparations was examined for the purpose of understanding the inactivation of free enzymes. The substance of the approach is in preventing to occur or modifying the activation energies of some reactions. Both effects contribute to revealing the reactions that were disguised at the analysis of free enzyme inactivation. A
80 unified evaluation procedure was tested on four preparations of invertase bound to the surface of functionalised cellulose beads. The enzyme coupling was strong enough to prevent dissociation and, consequently, association reactions during thermal inactivation.
1.0
0.8
I:! .. 0 . 6
~176176176176 ~176176176 ~176176 .... ~176176 ............
"=: 0 , 4
I
0.2
0.0
:- . . . . . --
Model Model
1.2 1.3
--"-'-
Mode!
2.6
I
I
,,I
I
I
I
1_
0
10
20
30
40
50
60
Time
(rain)
Figure 2. Comparison of the description of thermal inactivation of yeast invertase by tworeaction (1.2 and 1.3) and three-reaction (2.6) models. Reprinted from Enzyme Microb. Technol., Vol. 20, P. Vrfibel, M. Polakovi~, V. Stefuca and V. Bfile~, Analysis of mechanism and kinetics of thermal inactivation of enzymes: Evaluation of multitemperature data applied to inactivation of yeast invertase, 348-354, (1997), with permission from Elsevier Science. It was found that one of the principal criteria for assessing the suitability of immobilized preparations to complement the inactivation mechanism of free enzyme was the preservation of activity during immobilization. The biospecific adsorption on the concanavalin A-cellulose conjugate, giving the yield of immobilization of 74%, was clearly the most efficient technique in this respect. The high yield was achieved mainly because of the sparing effects of this immobilization technique on the active sites of invertase. 96% of the preserved activity, after subtracting the activity of unbound invertase, implied that this preparation could behave, in many respects, similarly to free enzyme. The excellent preservation of activity can be explained in that the invertase is bound here only through its carbohydrate moiety [9]. The superiority of the biospecific immobilization over the covalently bound enzyme or crosslinked preparations were confirmed also by the thermal stability and inactivation kinetics studies. The inactivation mechanism obtained was able to complement the previously verified mechanism of the inactivation of free invertase. Owing to prevention of dissociation and association reactions and decreased rates of inactivation, two more reactions were revealed which were not demonstrated at free invertase. The first one was in the denaturation sequence and the second one was assumed to modify the covalent bonds as it occurred in a significant measure only above 60~ [8].
81 4. MULTI-PH EVALUATION [10l The benefits of multiresponse evaluation has been extended also to the pH-induced inactivation since the effect of pH on the mechanism of enzyme inactivation can be well defined by a sequence of rapid protonation and deprotonation reactions [11,12]. Such an approach was tested on the inactivation of invertase at seven different pHs in the range of 2.19.2 and temperature of 20~ Again, the evaluation of single and multiple inactivation curves was compared. The former approach was proved to be completely unsuitable for the mechanism analysis since the pH-dependence of each integral parameter was expressed through five intrinsic parameters. On the other hand, the multi-pH evaluation provided a consistent model with statistically significant parameters. The verified model represented the following mechanism, ,
E r------~ D1
~K 1
k;
>D 1
~K1
E;(4)
(2)
2
E3
k, ;E'3[a'3)""
k', ; D 3
where both native enzyme and denaturation intermediate (labelled with dash) existed in three ionisation forms. El is the acidic form of native enzyme which dissociates into the neutral form E2 by liberating hydrogen ion and the deprotonation reaction is unambiguously characterised by the equilibrium constant Kl. The same applies also for the transition of E2 into the alkaline form E2 as well as for the ionisation reactions between the different ionisation states of active intermediates, D'1, E'2 and E'3 . Only the pool of the molar activities of the native enzyme forms was determined and the molar activities of the intermediate enzyme forms, a~ and a~, are related to this value. The molar activity of the acidic intermediate was evaluated to be equal to zero therefore the character D was used for its symbol as for the irreversibly inactivated forms of enzyme. In order to decrease the number of parameters, it was assumed that the equilibrium constants characterising the transition between the acidic and neutral forms as well as between the neutral and alkaline forms of both native and intermediate forms were identical. It was found that pKl = 2 and pK2 --- 7.5 which means that at optimum pH 4.6 invertase exist in a single ionisation form whose rate of inactivation at the temperature of 20~ was negligible (k2 = 0).
5. CONCLUSIONS It has been discussed the critical problem in the identification of the mechanism of enzyme inactivation which is to properly relate the phenomenon of the loss of enzyme activity with the changes occuring in the structure of enzyme molecule. The principal contradiction lies in that the activity is a single, integral quantity determined by the kinetics of corresponding catalytic
82 reaction whereas the structural changes can be manifold and their significance for the activity loss can be different. As the kinetics of enzyme activity loss gives the ultimate proof of the mechanism of enzyme inactivation, a rigorous quantitative analysis of experimental data is essential for the identification of true inactivation mechanism. It has been demonstrated that improved design and evaluation of inactivation experiments lead to the verification of more complex mechanisms than the conventional procedures. The principal contribution to improving the analysis of enzyme inactivation presented in this publication was the evaluation of multiresponse inactivation experiments. Main emphasis was on the procedures where a single process variable varied (temperature or pH, respectively). It has been shown that the variation of another factor could bring further benefits in identification of inactivation mechanism. It was a selective immobilization in this study; the variation of initial concentration of enzyme could be of interest in investigating the significance of bimolecular reactions in the inactivation pathway. The importance of the experimental techniques providing the information about structural changes in the protein molecules is not underestimated therewith. They have also a significant role in discrimination between alternative mechanisms. They, however, are of limited use, if they are not accompanied by a careful analysis of activity measurements. Moreover, these structural techniques provide mostly qualitative or semi-quantitative data in respect to the kinetics of enzyme inactivation. It would be topmost interesting to integrate quantitatively these types of data which means to evaluate simultaneously the inactivation data together with the results of monitoring of denaturation using calorimetric, spectroscopic or chromatographic techniques.
REFERENCES 1. 2. 3.
M. Polakovi~ and P. Vr~ibel, Process Biochem., 31 (1996) 787. V.V. Mozhaev and K. Martinek, Enzyme Microb. Technol., 4 (1982) 299. M. Dixon, E.C. Webb, C.J.R. Thome and K.F. Tipton, in Enzymes, 3rd ed., Longman, London, 1979, 165. 4. A. Sadana, Trends Biotechnol., 6 (1988) 84. 5. W.R. Lencki, J. Arul and R.J. Neufeld, Biotechnol. Bioeng., 40 (1992) 1421. 6. W.R. Lencki, J. Arul and R.J. Neufeld, Biotechnol. Bioeng., 40 (1992) 1427. 7. P. Vr~ibel, M. Polakovi~, V. ~tefuca and V. B~ile~, Enzyme Microb. Technol., 20 (1997) 348. 8. P. Vr~ibel, M. Polakovi~,, V. God6, V. B~ile~, P. Do~olomansk~, and P. Gemeiner, Enzyme Microb. Technol., 21 (1997) 196. 9. J. Turkowi, M. Fusek and J. ~t'ovi~kowi, in A. Bla~ej and J. Zemek (eds), Interbiotech '87. Enzyme Technologies, Progress in Biotechnology, Vol. 4, Elsevier, Amsterdam, 1988, 245. 10. P. Vr~ibel, Study on the Inactivation of Biocatalysts, Ph.D. thesis, Slovak Technical University, Bratislava, 1995. 11. K.J. Laidler, The Chemical Kinetics of Enzyme Actior~ Clarendon Press, Oxford, 1958. 12. A. Chitnis and A. Sadana, Biotechnol. Bioeng., 34 (1989) 804.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
83
Stability and stabilization of a - l , 4 - D - g l u c a n p h o s p h o r y l a s e s R. Gdegler a, B. Mtiller-Fembeck a, S. D'Auria b, F. La Cara b, and B. Nidetzky a a Division of Biochemical Engineering, Institute of Food Technology, Universittit fiir Bodenkultur Wien, Muthgasse 18, A-1190 Wien, Austria b Institute of Protein Biochemistry and Enzymology, Consiglio Nazionale delle Ricerche, via Marconi n. 10, 80125 Napoli, Italy
1. I N T R O D U C T I O N The a-l,4-D-glucan phosphorylases catalyze the reversible degradation of glycogen or other storage polysaccharides into ct-O-glucopyranosyl-1-phosphate (Glc-l-P) using pyridoxal 5'-phosphate (PLP) as the essential cofactor [1]. In addition to their key role in cellular carbohydrate metabolism, glucan phosphorylases are useful catalysts in carbohydrate synthesis by providing Glc-l-P as activated glucosyl donor [2]. Interactions of glucan phosphorylases with phosphoryl groups are involved in catalysis [1,3] as well as in regulation of enzyme activity [3] and could be even important determinants of conformational stability of phosphorylase. Multiple phosphate-binding sites have been located in classes of nonregulated and regulated ct-glucan phosphorylases [3] and involve interactions with the phosphate residue of the cofactor PLP, the substrate phosphate, the allosteric effectors AMP or glucose-6phosphate as well as covalently phosphorylated Ser or Thr residues [3]. These proteinphosphate interactions have been studied at a molecular level [3,4]. In spite of detailed investigations on the interactions of phosphorylase with the phosphate anion in relation to catalysis and enzyme activation, a role of phosphate or phosphorylated metabolites in stabilization of glucan phosphorylases to, e.g., thermal inactivation, has not received much attention. Here, we present data that support a stabilizing function of the phosphate anion with several members of the glucan phosphorylase family. The most remarkable effect was observed with nonregulated phosphorylases from, e.g., mesophilic bacterium Corynebacterium callunae or plant material such as potato tuber. In case of C. callunae phosphorylase, the interaction with phosphate enables this enzyme to acquire a resistance to thermal inactivation that is typical for enzymes from thermophilic rather than mesophilic sources. Regulated mammalian phosphorylase that contains the allostenc effector site displayed significant stabilization by AMP. The effect, however, was weak when compared with phosphate-dependent stabilization in bacterial or plant phosphorylases.
84 2. M E T H O D S
2.1 Giucan Phosphorylases Rabbit muscle phosphorylases a and b were the commercially available preparations from Bochringer (Mannheim, Germany). E. coil maltodextrin phosphorylase was produced in E. coli JM 109 harboring the plasmid pMAP101 (kindly provided by R. Schinzel, Wiirzburg, Germany) following induction with 0.2 % w/v maltose [5]. The enzyme was purified to homogeneity by hydroxyapatite chromatography as previously described [6]. Glucan phosphorylase from C. callunae was obtained as reported recently [7]. Starch phosphorylase from Solanum tuberosum was isolated following published procedures [8] with slight modifications, using hydrophobic interaction chromatography on Phcnylsepharose fast flow and subsequent ion exchange chromatography on DEAE-sepharose. A crude cell extract of thcrmophilic Thermusflavus AT 61 that contained glucan phosphorylase activity was used. 2.2.
Assays Phosphorylasc activity was determined at 30~ in phosphorolysis direction (50 mM
potassium phosphate, pH 6.9) employing a coupled assay with phosphoglucomutase and NAD+-dependent glucose-6-phosphate dehydrogenase [7]. Twenty g/L maltodextrin (Agrana, Austria) was used as glucan substrate. For measuring the activity of the muscle glycogen phosphorylase, 15 g/L oyster glycogen (Sigma, Germany) and 1 mM AMP were employed. The activity of the T. flavus enzyme was determined in a discontinuous enzymatic assay by measuring the Glc-I-P released from maltodextrin after 1 h incubation at 65~
2.3. Stability to inactivation by elevated temperature Following preselection of the suitable incubation temperature (40 - 65~
the
phosphorylases (= 1- 2 U/mL) were incubated in 50 mM Tris.HCl buffer, pH 8.0, in the absence or presence of phosphate, AMP or other phosphate analogs (20 - 50 mM each). For easier comparison, reactions that were designated phosphate-free contained 0.5 m M phosphate because traces of phosphate are difficult to remove by gel filtration and were found to affect the thermal stability of the bacterial phosphorylases significantly. Samples were taken in regular intervals from the incubation mixtures, and the residual enzyme activity was immediately determined. Inactivation constants were calculated assuming pseudo first-order decay of activity, and enzyme half lives were derived accordingly.
2.4. Stability to inactivation by proteolysis Phosphorylase (= 2 pM) was incubated at 30~ with Proteinase K (Boehfinger; 2 or 6 lag/mL) in the absence and presence of phosphate, AMP or analogs (50 mM each). Samples taken from the reaction mixture were immediately assayed for phosphorylase activity.
85 3. R E S U L T S
3.1. Nonregulated glucan phosphorylases The phosphorylases from E. coli, C. callunae and likely T. flavus are members of the class of nonregulated bacterial glucan phosphorylases that lack regulation of enzyme activity by allosteric effectors and covalent phosphorylation [3,4]. The plant enzymes, e.g., from S.
tuberosum (potato tuber) also belong to this class. When incubated at temperatures ranging from 40 to 65~
marked differences in stability to irreversible thermal inactivation were found for
these four glucan phosphorylases (Table 1). Plots of the natural logarithm of residual enzyme activity as function of incubation time were linear suggesting that first-order model of inactivation is applicable for data analysis. Based on thus calculated half-life of enzyme activity at 50~ in the absence of phosphate ions, the enzymes can be ranked according to increasing thermostability as C. callunae << E. coli < S. tuberosum << T. flavus. Not unexpectedly, the activity of phosphorylase from thermophilic T. flavus was constant during incubation at 65 ~ for 24 h (which was the maximum incubation time in the experiments), and the half life of the enzyme at this temperature was therefore not determined. Table 1 Stability to temperature-induced inactivation of nonregulated phosphorylases. i
ii
Half-life (min) i
No addition a
ii
ii
50 mM Phosphate
50 mM AMP
lllll
i
E. coli maltodextrin phosphorylase (45 ~
10
24
24
C. callunae starch phosphorylase (50 ~
=1
stable b
-- 1
S. tuberosum starch phosphorylase (50 ~
43
stable b
n.d.
T. flavus glucan phosphorylase (65 ~
stableb
stableb
n.d.
i
n.d., not determined; a 50 mMTris.HCl buffer, pH 8.0; b incubation time, 3 h. For the least thermostable phosphorylase (that from C. callunae) the buffer component was found to play a significant role in determining the stability to heat-induced inactivation. According to relative stabilization of enzyme activity at 30~
buffers (50 mM) were ranked as
acetate = glycerophosphate = triethanolamine > imidazole >> Mes = Tris. Compared with, e.g., glycerophosphate, imidazole had a clear destabilizing effect and, in concentrations of > I(X) m M
86 facilitated removal of PLP in C. callunae phosphorylase (not shown). In Tris- and Mesbuffered solutions phosphorylase from C. callunae lost about 50% of its activity at 4~ within 12 h. In the presence of 50 mM phosphate, however, the picture of relative thermostability was changed drastically (Table 1). With the exception of thermostable phosphorylase from T. flavus for which no inactivation was observed at 65 ~ anyway, significant stabilization by phosphate was observed for all other phosphorylases. For the enzyme from E. coli the stabilizing effect was only moderate, giving an approximately 2.5-fold increase in stability at 45 ~ compared with the control lacking the phosphate. In contrast, for phosphorylases from S. tuberosum and C. callunae the competence of phosphate to confer extra stability was very high. In the presence
of 50 mM phosphate no inactivation was seen at 50~
with both enzymes for at least 3 h.
Taking into consideration the moderate stability of the enzyme from C. callunae at this temperature in the absence of phosphate ion, the resistance of this phosphorylase to inactivation in phosphate buffer is remarkable. Even at 60 ~
no loss of enzyme activity was detected after
an incubation time of 1 h, indicating that protein-phosph~fte interactions are capable of conferring stability properties to this enzyme that are typical for thermophilic proteins. The apparent specificity of the stabilizing interaction is interesting. Structural analogs of phosphate such as sulfate or arsenate could confer extra stability to C. callunae phosphorylase at 50~ although with at least 10-fold reduced efficiency relative to phosphate. Phosphate analogs ( 150 mM) such as phosphite, thiophosphate or fluorophosphate or the substrate Glc-I-P (50 mM) did neither stabilize by themselves nor inhibit the stabilizing effect by 50 mM phosphate at 50 ~
Nonregulated phosphorylases are thought to contain a primitive form of the aUosteric AMP
site found in regulated mammalian glycogen phosphorylase. It is therefore interesting to note that AMP (50 mM) stabilized phosphorylase from E. coli equally well as phosphate (50 mM). On the other hand, AMP was completely ineffective to stabilize the phosphorylase from C. callunae.
3.2. Regulated glucan phosphorylase In contrast to their nonregulated counterparts, regulated phosphorylases such as rabbit muscle glycogen phosphorylase contain the highly developed allosteric effector site (which involves interaction with phosphorylated components) and the covalent phosphorylation site. Hence, the regulated phosphorylases contain at least one additional phosphate site which could potentially participate in stabilization of the protein. In the absence of phosphate or AMP phosphorylase a was less stable than the unphosphorylated b-form of the enzyme (Table 2). Phosphate ions could not stabilize glycogen phosphorylase a and b to thermal inactivation, but AMP did. The effect was slightly higher for the phosphorylated a -form of the enzyme for which an about 30-fold stabilization of enzyme activity by the nucleotide was found. Compared with the stability of nonrcgulatcd phosphorylascs from C. callunae and S. tuberosum in the
87 presence of 50 mM phosphate, the resistance of glycogen phosphorylase to heat inactivation at 50 ~ even with added AMP was quite low. Table 2 Stability to temperature-induced inactivation of regulated phosphorylase. Half-life at 50 "C (min) ii
No addition a
50 mM Phosphate
20mM AMP
Rabbit muscle glycogen phosphorylase b
17
16
53
Rabbit muscle glycogen phosphor),lase a
4
5
117
ii
a 50 mMTris-HCl buffer, pH 8.0. 3.3 S t a b i l i z a t i o n
to proteolysis
The increased conformational stability of glucan phosphorylases in the presence of phosphate or AMP, compared with the control that lacked these components, was demonstrated further by measuring the loss of enzyme activity as function of incubation time at 30 ~ when proteinase K was added. The results in Table 3 corroborate what has been found for temperature-induced inactivation (Tables 1 and 2). Again, there was a pronounced stabilizing effect by phosphate with the starch phosphorylase from C. callunae that was fully resistant to proteolytic attack in the presence of 50 mM phosphate. On the other hand, AMP stabilized rabbit muscle phosphorylase and E. coli maltodextrin phosphorylase, but not the enzyme from C. callunae.
Table 3 Stability to proteinase-induced inactivation of nonregulated and regulated phosphorylases i
i
Half-life (min)
i
i
No addition a
50 mM Phosphate
50 mM AMP
E. coli maltodextrin phosphorylase b
22
60
65
C. callunae starch phosphorylase b
110
stable d
100
Rabbit muscle glycogen phosphorylase b c
19
18
30
Rabbit muscle glycogen phosphor),lase a c
11
11
23
a 50 mM Tris.HC1 buffer, pH 6.9; b 6.0 ktg/mL and c 2.0 I.tg/mL proteinase K; d 3 h.
88 4. D I S C U S S I O N The results described here show that interactions with phosphate not only play key roles in catalysis and regulation of activity [3,4], but also in determining the conformational stability of several ix-1,4-D-glucan phosphorylases. Many proteins are known to interact with phosphate or phosphorylated components, and the phosphate ion has been previously shown to confer extra stability to other enzymes such as aspartate aminotransferase [9] or RNAse [ 10]. Most protein-phosphate interactions are thought to occur between (i) the phosphate oxygens and the main chain nitrogens at the start of an ~t-helix or (ii) with arginine residues in nonhelix interactions [ 11]. The common phosphate recognition sites found in regulated and nonregulated phosphorylases comprise the active site, where phosphate or Glc-I-P bind, and a positively charged arginine cluster that is responsible for binding of the phosphoryl group of AMP or glucose-6-phosphate at the allosteric effector/inhibitor site of glycogen phosphorylase 13,4]. Interestingly, apart from the active-site phosphate a second bound phosphate anion was located in the recently solved crystal structure of E. coli maltodextrin phosphorylase [ 12]. This second phosphate was found to interact at a similar position to that where AMP binds in rabbit muscle glycogen phosphorylase. In conclusion, it seems quite interesting to probe, e.g., by chemical modification and/or site directed mutagenesis, the involvement of different residues in glucan phosphorylase stability and stabilization by phosphate in more detail. Pertaining to use of phosphorylases in carbohydrate synthesis, stabilization by phosphate could certainly be important in order to improve utilization of enzyme activity in conversion of a-glucans into Glc1-P or synthesis of amylose chains. REFERENCES
*
3. 4. 0
0
0
o
9. 10. 11. 12.
D. Palm, H. W. Klein, R. Schinzel, M. Buehner and E. J. M. Helmreich. Biochemistry, 29 (1990) 1099. G. Ziegast and B. Pfannemiiller, Carbohydr. Res., 160 (1987) 185. L. Johnson, FASEB J., 6 (1992) 2278. C. B. Newgard, P. K. Hwang and R. J. Fletterick, CRC Crit. Rev. Biochem. Mol. Biol., 21 (1987) 69. A. Weinh~iusel, B. Nidetzky, C. Kysela, and K. D. Kulbe, Enzyme Microb. Technol., 17 (1995) 140. C. Eis, R. Griel]ler, A. Weinh~usel, B. BSck, K. D. Kulbe, D. Haltrich, R. Schinzel and B. Nidetzky, J. Biotechnol., 58 (1997) 157. A. Weinh~iusel, R. Grietler, A. Krebs, P. Zipper, D. Haltrich, K. D. Kulbe and B. Nidetzky, Biochem. J., 326 (1997) 773. J. Hollo, E. Laszlo and A. Hoschke, Starch/St~ke, 16 (1964) 243. J. H. Martinez-Liarte, A. Iriarte and M. Martinez-Carrion, Biochemistry, 31 (1992) 2712. E. M. Meiering, M. Bycroft and A. R. Fersht, Biochemistry, 30 (1991) 11348. P. Chakrabarti, J. Mol. Biol., 234 (1993) 463. K. Watson, R. Schinzel, D. Palm and L. N. Johnson, EMBO J., 16 (1997) 1.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
89
Oxidation by h y d r o g e n peroxide o f D - a m i n o acid oxidase from Rhodotorula
gracilis V. Obreg6n, I. de la Mata, F. Ram6n, C. Acebal and M.P. Castillrn Biochemistry and Molecular Biology I Department, Faculty of Biology, Universidad Complutense, 28040 Madrid, Spain.
D-amino acid oxidase (DAAO) from Rhodotorula gracilis is a flavoenzyme that catalyzes the oxidative deamination of D-amino acid to give the corresponding tx-keto acids, hydrogen peroxide and ammonia. This enzyme exhibits interesting properties that enable its use in industrial production of semisynthetic cephalosporins. Hydrogen peroxide induced enzyme deterioration that was more remarkable in the case of the apoenzyme when compared with the native holoenzyme. The inactivation rate constant was 3.12xl 02mM~min ~ for the apoenzyme and 6.96~10SmMlmin-1 for the holoenzyme. Fluorescence emission studies have shown the oxidation of tryptophan residues, by H202, as responsible of enzyme inactivation. The oxidized tryptophans did not seem to be involved in the coenzyme FAD binding to DAAO apoprotein.
1. INTRODUCTION D-amino acid oxidase (EC 1.4.3.3, DAAO) is a flavoenzyme that catalyzes the oxidative deamination of D-amino acids to give the corresponding ct-keto acids and ammonia. Native holoenzyme from Rhodotorula gracilis is a 80 kDa dimer of identical subunits containing a molecule of tight binding FAD per monomer. FAD, which is reduced during the course of the reaction, is further reoxidized by 02 with the production of hydrogen peroxide (1, 2). Damino acid oxidase was isolated and purified from different sources by far (2), but the enzyme from yeast Rhodotorula gracilis is very interesting for biotechnological purposes due to its high turnover, tight binding to FAD and high activity on eephalosporin C (3). D-amino acid oxidase from Rhodotorula gracilis efficiently oxidizes cephalosporin C to otketo adipil-7-ACA which is further non-enzymatically decarboxylated to glutaryl-7-ACA which can be hydrolyzed by glutaryl-7-ACA acylase to 7-ACA, an important starting material to obtain semisynthetic cephalosporins (4, 5). Hydrogen peroxide is a by-product in the oxidation of D-amino acids by D-amino acid oxidase that is important for the non-enzymatic decarboxylation of the keto-adipic moiety of ct-keto-adipil-7-ACA; however, it can induce enzyme deterioration in industrial bioreactors (5, 6). Hydrogen peroxide is a potent oxidant for many organic compounds. It was found that under relatively mild conditions this reagent is highly specific for a small number of amino acids side chains (7) including thioether, indol, sulfhydryl, disulfide, imidazole and phenolic.
90 Under acidic conditions the primary reaction is the conversion of methionine residues to the corresponding sulfoxide. Oxidation of cystine or cysteine is favored by increased pH and the presence of heavy metals in the case of cysteine. Tryptophan and tyrosine residues in proteins are more resistant to attack but tryptophan oxidation occurs optimally between pH 8 and 10. Although free histidine is attacked under the same conditions it appeared to be resistant when present in polypeptide linkage. In the present study we report the oxidation of tryptophan residues in D-amino acid oxidase from Rhodotorula gracilis as the responsible of enzyme inactivation in the presence of hydrogen peroxide.
2. MATERIALS AND METHODS
2.1. Materials D-alanine, hydrogen peroxide, pyruvic acid, FAD and 2,4-dinitrophenylhydrazine were from Sigma Chemicals (St. Louis, MO, USA). All other reagents solvents were of analytical grade and purchased from Merck (Darmstadt, Germany). 2.2. Enzyme purification D-amino acid oxidase was isolated and purified from Rhodotorula gracilis (American Type Culture Collection, strain number 26217) cultures as previously described (8). Apoenzyme was prepared by dialysis against potassium bromide, as described by Casalin et al. (9), using a strategy originally proposed for preparation of mammalian apo-D-amino acid oxidase (10). 2.3. Enzyme assay The activity of D-amino acid oxidase was determined at 35~ in 1001al air saturated incubation mixtures containing 87.5ng of enzyme and 10mM D-alanine in 50mM potassium phosphate buffer at 8.5. The released pyruvic acid was determined after 10 minutes by reacting with 2,4-dinitrophenylhydrazine and the corresponding hydrazone was monitored at 450nm. In inactivation experiments, H202 was removed from the mixtures by adding catalase in the assay buffer. Concentration of H202 in stock solutions was checked spectrophotometrically by using an extinction coefficient of 43.6 M tern ~ at 240nm. The absence of oxidative decarboxylation by H202 of the r formed was checked by incubation of 250~tM pyruvate solution with 5 and 50mM H202 at room temperature and comparison of the spectra of the corresponding 2,4-dinitrophenylhydrazones. 2.4. Fluorescence spectra Fluorescence spectra for both apo- and holo-D-amino acid oxidase before and after oxidation with hydrogen peroxide were recorded in a MPF-44E Perkin-Elmer spectrofluorimeter. 1 ~tM apo- and holoenzyme solutions were oxidized with 10ram H202 for 30 minutes and 50raM H202 for 12 hours respectively, at 30~ A reference mixture was prepared for each oxidation reaction in which H202 was substituted by H20. After incubation, H202 was removed by using columns of semi-dry Sephadex G-25. Then, emission spectra were recorded at room temperature.
91 2.5. Characterization of binding of FAD to oxidized apo-enzyme Binding of FAD to oxidized apo-D-amino acid oxidase was followed by recording FAD fluorescence emission. Upon excitation at 450nm, the fluorescence emission was recorded at 530nm (11). Dissociation constant of FAD enzyme complex was calculated by fitting the experimental data in figure 3 to the following equation (12) (1)
1/(l-a) = (1/Kd [FAD/a]) - (l~(d [DAAO])
where "a" is the fraction of the total FAD-D-amino acid oxidase binding sites and Kd is the dissociation constant of FAD-D-amino acid oxidase complex.
3. RESULTS AND DISCUSSION The apoenzyme of Rhodotorula gracilis was obtained with a good yield. It is a 40kDa monomer under all conditions (9). D-amino acid oxidase (holo- and apoenzyme) was preincubated with 1-50mM hydrogen peroxide at the indicated times in Materials and Methods section. Holoenzyme was rather resistant to the deleterious effect of the reagent since its activity was unchanged atter 12 hours of preincubation with 1-5mM of hydrogen peroxide. The inactivation followed pseudo-first order kinetics wiht a second-order rate constant of 6.96x10SmMlminl. By contrast the apoenzyme was very sensitive to hydrogen peroxide with a second-order rate constant of inactivation of 3.2xl 0"2mMlmin 1, three orders of magnitude higher than for the holoenzyme (Figure 1).
120 I00
9-
80
ol--t +.a
o
60
~
4o
<
20 0
30
60 Time
90
120
150
180
210
(minutes)
Figure 1. Kinetics of inactivation of DAAO by H202. Apoenzyme preincubated with: l mM H202 (-e-), 10mM H202 (-V-). Holoenzyme preincubated with: l mM H202 (-II-), 10mM H202 (-0-), 50mM H202 (-A-).
92 Since it has been shown that the oxidation of tryptophan residues by hydrogen peroxide occurs optimally between pH 8 and 10 (13), titulation of tryptophans of holo and apoenzyme before and after treatment with the reagent has been carried out. The fluorescence emission spectra for the different forms of the enzyme are shown in figure 2.
1
FI
FI F
2~5
' nm
45'0
I
290
.....
nm
[
450
Figure 2: (A) Fluorescence emission spectra for the native apoenzyme (1, gain 3), and for the H202-treated apoenzyme (2, gain 10). (B) Fluorescence emission spectra for the native holoenzyme (1, gain 10) and for the H202-treated holoenzyme (2, gain 10).
There was an important change in the fluorescence emission between the apoenzyme and the H202-treated apoenzyme indicating that hydrogen peroxide oxidized tryptophans in the apoenzyme (Figure 2A). Spectra for the holoenzyme and H202-treated holoenzyme are more similar due to less tryptophans are exposed to the surrounding medium in the holoenzyme because of FAD binding and dimerization. On the other hand,when comparing the spectra for apo- and holo-enzyme it can be observed a considerably higher fluorescence intensity for the native apoenzyme than for the native holoenzyme, probably due to the quenching of tryptophans when FAD binds to the apoenzyme. Finally, native holoenzyme shows a more structured spectra than apoenzyme does (Figures 2A and 2B) with two maximums at 325 and 335nm, indicating the presence of two tryptophan families that belong to two structural different domains in the protein. Oxidized enzyme binds FAD with a dissociation constant of 0.5x10"aM (Figure 3) that is similar to that determined for the native holoenzyme (9). This result shows that the oxidized tryptophans are not in the FAD-binding site.
93 350
250
FI 150
50 0,1
0,2
0,3
0,4
[FAD] ( ~ M ) Figure 3: Titulation of H202-treated apoenzyme with FAD. The change of fluorescence emission was monitored at 530nm.
Previous studies with chemical modifiers allowed to have an insight into the active site of the yeast enzyme. To date, histidine, lysine, arginine and cysteine have been described as competent for the oxidation of D-amino acid oxidase from Rhodotorula gracilis (8, 14-16); however, this paper presents the first evidence about the possible relevance of tryptophan residues in either substrate binding and catalysis or in maintaining the enzyme in catalytically competent conformation. Studies are in progress to attempt discriminate between these possibilities.
REFERENCES 1. M. Pilone Simonetta, L. Pollegioni, P. Casalin, B. Curti and S. Ronchi, Eur. J. Biochem., 180 (1989) 199. 2. B. Curti, S. Ronchi and M. Pilone Simonetta, F. Muller (ed) Chemistry and Biochemistry ofFlavoenzymes, Vol. III, pp. 69-94, CRC Press Boca Raton, FL, 1992. 3. L. Pollegioni, A. Falbo and M. Pilone Simonetta, Biochim. Biophys. Acta, 1120 (1992) 11. 4. T.A. Savidge, EJ. Vandamme (ed) Biotechnology of Industrial Antibiotics, pp.205-215, Dekker NY, 1984. 5. E. Szwajeer Dey, S. Flygare and K. Mosbach, Appl. Biochem. Biotechnol., 27 (1991) 239. 6. M. Pilone Simonetta, L. Pollegioni and S. Buto, Biotechnol. Appl. Biochem., 16 (1992) 252. 7. N.P. Newman, Methods" Enzymol., 25 (1972) 393. 8. F. Ram6n, I. de la Mata, S. lannacone, M.P. Castill6n and C. Acebal, J. Biochem., 118 (1995)911. 9. P.Casalin, L. Pollegioni, B. Curti and M. Pilone Simonetta, Eur. J. Biochem., 197 (1991) 513. 10. V. Massey and B. Curti, J. Biol. Chem., 241 (1966) 3417.
94 11. S. Ghisla, V. Massey, J.M. Lhoste and S.G. Mayhew, Biochemistry, 13 (1974) 1. 12. R.A. Stinson and J.J. Holbrook, Biochem..L, 131 (1973) 719. 13. Y. Hachimori, H. Hrinishi, K. Kurihara and K. Shibata, Biochim. Biophys. Acta, 93 (1964) 346. 14. G. Gadda, G.L. Beretta and M. Pilone Simonetta, Biochem. Mol. Biol. Int., 33 (1994) 947. 15. G. Gadda, A. Negri and M. Pilone Simonetta, J. Biol. Chem., 269 (1994) 17809. 16. L. Pollegioni, S. Campaner, A.A. Raibekas and M. Pilone Simonetta, Arch. Biochem. Biophys., 343 (1997) 1.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
95
Cosolvent effect on the synthesis o f ampicillin and cephalexin with penicillin acylase C. Aguirre a, J. Baeza a and A. Illanes b. ILaboratofio de Recursos Renovables, Universidad de Concepci6n. PO Box 20-C, Concepci6n, Chile bEscuela de Ingenieria Bioquimica, Universidad Cat61ica de Valparaiso. PO BOX 4059, Valparaiso, Chile
ABSTRACT Penicillin acylase produced from Bacillus megaterium ATCC 14945 and a commercial immobilized enzyme from Escherichia coil were studied for the syntheses of ampicillin and cephalexin under kinetic control, considering the effect of dimethyl sulfoxide (DMSO), methanol and sorbitol as organic cosolvents. Ampicillin and cephalexin yields were higher for the B. megaterium enzyme: 21.3 % and 45.5% respectively, as compared to 14.5 and 29.5% for the enzyme fTom E. coll. Sorbitol (20% w/v) increased ampicillin and cephalexin yields by 28 an 20% respectively and enzyme activity was completely recovered in this reaction medium after 12 h. By contrast, yields were lower in DMSO (20% v/v) and methanol (20% v/v), enzyme suffering inactivation in both media.
1. INTRODUCTION Penicillin acylasc (PA) has been used traditionally in the hydrolysis of penicillin G and cephalosporin to produce 6-aminoponicillanic acid (6-APA) and 7-aminocephalosporanic acid (7-ACA), precursors of the semisynthetic penicillins and cephalosporins respectively [1 ]. PA can be used, however, as a synthetase to produce 13-1actam antibiotics from such nuclei. Syntheses can be carded out by direct condensation of the [3-1aetam nuclei (nucloophile) with the acyl donor (thermodynamic or equilibrium-controlled synthesis) or by reaction of the nucleophiles with an activated acyl donor, such as an ester or amide of the respective acid (kinetic-controlled synthesis). In the former case, yield will depend of the thermodynamic equilibrium; in the latter, yield will result from the balance of three different enzymatic reaction rates: synthesis of antibiotic (Vs,A), hydrolysis of the already synthesized antibiotic (Vh,A) and hydrolysis of the activated acyl donor (Vh,B). Nam et al. [2] and Kasche et al. [3],
96 postdated the formation of an acyl-enzyme intermediate which is either attacked by the nucleophile to synthesize the antibiotic or attacked by water to promote hydrolysis. The syntheses of ampicillin and cephalexin under kinetic control have been studied, using PA from E.coli and B.megaterium. Effect of cosolvents (DMSO, methanol and sorbitol) on antibiotic yield is reported. Equilibrium-controlled synthesis was not considered owing to the low solubility of the substrates required for the syntheses.
2. METHODOLOGY PA from B.megaterium was produced by fed-batch fermentation in casein-hydrolyzate medium with phenylacetic acid as inducer [4]. Immobilized PA from E. coli was a commercial product from Boehringer-Mannheim, Germany. Syntheses of arnpicillin and cephalexin were performed at 37~ pH 7.0, 0.5 IU PA/mL, 20 mM 6-APA (or 7-aminodeacetoxycephalosporanic acid: 7-ADCA) and 60 mM phenylglycine methyl ester (PGME). Reaction kinetics were determined by HPLC, using a Lichrospher 100 RP-18 column (125 x 4 mm, 5 gin) from Merck (Darmstadt, Germany). Samples were eluted isocratically with 70% (v/v) 20 mM phosphate buffer pH 6.0 in methanol at a flow rate of 0.9 mL/min. Absorbance was recorded at 220 nm. Enzyme stability was determined as residual (hydrolytic) activity after 12 hours. Activity was determined by the Balasingham method [5] with penicillin G potassium salt (PGK) as substrate. One international unit of PA 0U) is the amount of enzymes releasing one grnol of 6APA per minute from PGK at pH 7.0 and 37~
3. RESULTS Kinetics of syntheses of ampicillin and cephalexin with both enzymes are presented in Figure 1. Yield maxima are observed, as expected from a kinetieally controlled reaction, yield values being reduced as product hydrolysis outweighs synthesis. Yields on ampicillin and cephalexin were higher with B. megaterium PA, as seen in Table 1. Product hydrolysis rates were higher with the E. coli enzyme, therefore reducing yield. Yields and initial rates of synthesis of eephalexin were always higher than ampicillin (Table 1), 7-ADCA being a better nucleophile than 6-APA for both enzymes.
Table 1 Yields (YA,Yc) and initial rates of synthesis (vs,A, Vs,c) of ampicillin (A) and cephalexin (C). i
Enzyme B. megaterium E. coli
YA
Yc
(%)
(%)
(mM )
(mS )
21.3 14.5
45.5 29.5
1.91 2.73
8.86 7.84
Vs,A
Vs,C
97 100
200
300
10
40
t,.-
Q.
r
(.)
500 50
E8 t~
400
e-.
6
30 .~
4
ao g
ID >
o 10
51-
-125
~. 9
10 0~
<
5 0
Time (min)
Figure 1. Kinetics of synthesis of cephalexin (A) and ampicillin (B) with PA from
B.megaterium (A) and E.coli ~).
Effect of cosolvents at 20% (v/v) is shown in Figure 2 for the synthesis of ampicillin with B.
megaterium PA. As seen in Table 2, sorbitol increased both yield (by 28%) and initial rate (2.4 times) of synthesis of ampicillin, while the opposite occurred with methanol and DMSO. Ratio of synthesis to hydrolysis rates (Vs,A/Vh,A) was higher with sorbitol and DMSO and lower with methanol. As seen in Table 2, PA was completely stable in sorbitol, while 10% and 22% inactivation occurred in the presence of DMSO and methanol respectively. This explains why yield on DMSO is lower even though Vs,A/Vh,Awas higher.
98 f
'
I
'
I
'
I
'
I
'
I
'
I
6
/
-~ 30
5
-
- 25~
v
2O ~ ~
-9
.o_. <E
~
-
1
3
-9
-
9
9
"
1H,,~ f
0
.
o
150
-15
100
200
300
400
500
600
Time / min
Figure 2. Effect of cosolvents (20% v/v) on the synthesis of ampicillin with B. megaterium PA. (V): methanol; (O): DMSO; 01): control; (&): sorbitol (20% w/v).
Table 2. Cosolvent effect on yields (YA) and initial rate (Vs,A) of ampicillin synthesis with B. megaterium PA.
Cosolvent
Control Sorbitol DMSO Methanol
........ YA
,,
Vs,A
(%)
(raM/h)
21.3 27.6 13.8 3.4
1.91 4.5 0.87 0.33
,
"
,
,
' Vs,A/Vh,A
01141 0.219 0.218 0.088
Stability (% activity after 12 hours ) 100 100 90 78
99 As shown in Figure 3, 20% (w/v) sorbitol also increased cephalexin yield, by 20%, an effect already reported for the E. coli PA [6]. Higher yields with sorbitol can be explained by the reduction in water activity that depresses hydrolytic reactions, and by its stabilizing effect on the enzyme. 12
60
10
50 A
~-,
8
40~
co
E v .E X
L_
30 > C
6
0
(3
m r e~
G)
(3
4
20
10
0
100
200
Time
300
400
500
(min)
Figure 3. Effect of sorbitol on the synthesis of cephalexin with B. megaterium PA. (1): control; (0):20% (w/v) sorbitol
4. CONCLUSIONS Better results were obtained with PA from B. megaterium both for the synthesis of ampicillin and cephalexin. 7-ADCA was a better nucleophile than 6-APA, higher yields being obtained in the production of cephalexin than ampicillin with both PA. Sorbitol increased yield and rate of synthesis of both antibiotics. However, the reported yields are still low to compete with existing chemical synthesis, despite the extremely mild conditions required for the enzymatic synthesis. Preliminary results indicate that pH, enzyme to substrate ratio and substrate to substrate ratio are important process variables, so that ample room exists for process optimization PA from B. megaterium is worthwhile to be studied in other organic solvent systems that produce no enzyme inactivation.
I00 REFERENCES 1. 2. 3. 4.
J. Shewale and V. Sudhakaran, Enzyme Microb. Technol., 20 (1997) 402. D. Nam, C. Kim and D. Ryu, Biotechnol. Bioeng.27 (1985) 953. V. Kasche, U. Haufler and L. Riechmann, Methods in Enzymol. 136 (1987) 280. A. IUanes, R. Tortes, O. Cartagena, A. Ruiz and M. Vdsquez, Process Biochem. 29 (1993) 263. 5. K. Balashingham, D. Warburton, P. Dunnill and M.D. Lilly, Biotechnol. Bioeng. 276 (1972) 250. 6. C. Hyun and J. Kim, Biotechnol. Bioeng. 42 (1993) 800.
Acknowledgements This work was supported by chilean FONDECYT, Grant N~ The authors express their gratitude to SINQUISA S.A., Lima, Peru for their kind supply of 6-APA and penicillin GK.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
101
Conversion and stability studies on enzyme-membrane reactor with lipase immobilized by different methods T. Venyige, E. Cs~myi and Cs. Sisak Research Institute of Chemical and Process Engineering, Pannon University of Agricultural Sciences, Egyetem u. 2,. H-8200 Veszpr6m, Hungary
1. INTRODUCTION Nowadays the enzyme-membrane reactors (EMR) have proved as a suitable system to carry out bioprocesses in which the components taking part in the reaction are in two immiscible liquid phases and the biocatalyst can be immobilized on or inside the membrane. They are favourably applied e.g. for enzymatic hydrolyses [1-3] and esterification [4,5] but there are several examples for other reactions, e.g. peroxidation of fatty acids [3]. The importance of membrane containing reactors is increasing further with the spreading of integrated systems. To perform simultaneously the biocatalytic reaction and the product removal it is obvious to utilize the opportunities of the membrane separation. Among the so-called extractive bioconversions, the first results about the formation of bioflavours was published ten years ago. The procedure was based on the finding that natural ethyl esters can be formed by in situ enzymatic esterification of fatty acids and ethanol produced from glucose via fermentation by yeast cells [6,7]. On the basis of the recent investigations [5], it has been cleared up that EMR is also applicable to ethyl oleate production if the ethanol content of substrate is of 10-30 wt%. *In the present studies, the performance of an enzyme-membrane reactor applied to ethyl oleate production is discussed. The main question to be answered is, how the reactor operates - concerning the ester formation and stability - under the circumstances of the really integrated extractive biocatalytic reactor system: i.e. at low ethanol content and in presence of living yeast cells. 2. MATERIALS AND METHODS 2.1 Materials Mucor michel lipase (LYPOZYME, 10000 LU/g, NOVO Nordisk Denmark) was used as catalyst. Aqueous ethanol (REANAL Hungary) solution of 60-160 g dm 3 initial concentration or ethanol containing fermentation broth produced by Saccharomyces bayanus
The financial supports of European Commission through COPERNICUS Grant No ERBCIPACT 923011 and German-Hungarian Intergovernmental Co-operation through Grant No. D/46-96 are gratefully acknowledged. " Acknowledgement:
102 IST 154 cells, and oleic acid (REANAL Hungary) were applied as substrates. Chemicals (from REANAL and Sigma Chemical Company) were of reagent grade.
2.2. Immobilization methods
Ultrafiltration/adsorption method: Lipase solution (100 cm 3, l g) was ultrafiltered (flow rate 4 cm 3 min -~) through the membrane and enzyme was immobilized by adsorption on the inner wall of the fibres. The process was followed by determination of protein contents of both the initial solution and permeate. The ratio between the amounts of dissolved and immobilized lipase was 3:2. The strength of the enzyme layer was checked by flowing distilled water through the membrane tubes: no protein wash-out was found [2]. Coating method: The undiluted enzyme solution (1 cm 3) was spread on the inner wall of the dry fibres applying alternated spreading and drying steps. After last drying, the fibres were coated with hexanoic solution of silicone rubber (DOW CORNING 3140 RTV, ingredients: methyl-trimetoxysiloxane, trimethylated silica, polymethylsiloxane). The 20 cm 3 solution contained 3 g rubber. In order to harden the silicone coating, the module was rinsed with inert gas for 24 hours. Since the polymerisation of silicone rubber takes place in presence of water, the gas stream was moistened. Emulsion method: The undiluted enzyme solution was mixed with the silicone solution (see above) and shaken in a test tube for 5 minutes. The emulsion was spread on the inner wall of the dry fibres as it was described above, with alternated spreading and drying steps. The drying procedure was the same as in case of the coating method. 2.3. Reactor configuration Experiments were carried out in a hollow fiber membrane bioreactor with 0.0188-0.0376 m 2 surface area. The hydrophylic membrane tubes ( asymmetric cellulose acetate, cut off: 40 000 Da) had 2 mm diameter and 200 mm length. During the reaction, 300 cm 3 oleic acid was recirculated on the enzyme side of the membrane and 100 cm 3 aqueous ethanol solution on the other side. Water produced could permeate through the membrane into the aqueous phase and ester remained in the organic phase. Concerning the studies in the extractive bioconversion system, the ethanol fermentation process was performed in a fluidized-bed immobilized-cell fermenter and the esterification in EMR described above. The scheme of the integrated system is illustrated in Fig. 1.
Figure 1. The scheme of the integrated fermenter/enzyme reactor system
103
Saccharomyces bayanus cells were immobilized by Ca-alginate gel entrapment. The fluidized-bed reactor was run in semicontinuous regime: periodically glucose solution of 40 wt % was added to the reaction mixture and the same quantity of broth was taken off. The effective volume of the reactor was 300 cm 3, its diameter 36 mm, the volume of the bed of immobilized-cell particles was 50 c m 3. The fermentation broth was recirculated in the primary side of the membrane reactor. 2.4. Analytical methods The samples were analyzed by HPLC (Merck-Hitachi L-6000A pump, Merck refractive index detector RI-71, temperature 70~ Nucleosil 100-10 C 18, 250x4 mm column with metanol:water (60:40) as eluent (flow rate 0,8 cm3/min) for acid and ester, PL Hi-PLex 300x7.7 mm column 5 mmol dm 3 H2SO 4 solution for ethanol (flow rate 0,7 cm3/min) were used.
3. RESULTS AND DISCUSSION
3.1. Esterification in enzyme-membrane reactors with iipase immmobilized by different methods Fig. 2 shows the bioconversion run in enzyme-membrane reactor with lipase immobilized by the ultrafiltration/adsorption method. The membrane surface was 0.0376 m 2. It can be seen on the figure that the rate of ester formation at the repeated adding of ethanol (dotted line) approximates the initial one, i.e. no considerable inactivation of enzyme can be supposed during the period.
1,6 1,4
:~~t........~
-
1,2 ~
Et(OH)
1
Ester
~ 0,8
~ 0,6 ~ 0,4
o,2 0 0
1
I
t
2
3
Time (days)
4
Figure 2. Time course of esterification in enzyme-membrane reactor with lipase immobilized by ultrafiltration/adsorption method. Time course of esterification applying the coating method is illustrated in Fig. 3. A model medium with non metabolizing Saccharomyces bayanus cells was used as aqueous phase (initial ethanol cone.: 163 mg/g, glucose cone.: 24 mg/g). Other parameters were the same as above. Comparing the initial slopes of the ester concentration vs. time curves in Figs. 2 and 3, it is unambiguous that the productivity of the enzyme-membrane reactor is higher in case of the coating method.
104 The experimental conditions applied to study the productivity of EMR with lipase immobilized by emulsion method were almost the same as that given previously, with exceptions of the enzyme fixation method and the volume of the organic phase (200 cm3). On the basis of the results (see Fig. 4), it is proved that some improvement in the productivity can be achieved using the emulsion method in comparison to the two other ones.
1,4
1
g
m
+__...._..-4
0,6 O
lira
0,4
Et(OH)
-
f
0,2
--a- Ester
0 .
I
I
I
t
I
0
0,2
0,4
0,6
0,8
1
1,2
Time (days) Figure 3. Time course of esterification in enzyme-membrane reactor with lipase immobilized by coating method.
1,6t
1,4~1,2-
" Et(OH) -.m-Ester
/~
1 0,8 I
0,6
elm
-a-t -a-t - - " " A
i
O
;~ 0,4
t
0,2 0 "~. 0
I
t
t
1
2
3
Time (days) Figure 4. Time course of esterification in enzyme-membrane reactor with lipase immobilized by emulsion method. 3.2. Extractive bioconversion in integrated system The organic phase of 200 c m 3 volume was recycled in enzyme-membrane reactor with 0.0188 m 2 membrane surface area. The time course of the run (Fig. 5) shows that quasi stationary conditions exist concerning ethanol concentration along a six day range. The ester level was increasing with almost constant slope, what implies probably that the equilibrium concentration would be much higher.
105
~0~ ~0,6
:. Et(OH) Ester e Glucose
O
~0,4 'r
t~--e i
0,2 __ O=9 0
, e------.e-....e.., e -
t~.
2
--e-.e" I
4 T ~ e (days)
,
I
ii
6
8
Figure 5. Time course of extractive bioconversion in fluidized-bed immobilized cell reactor/enzyme membrane reactor system.
3.3. Working stability of the enzyme-membrane reactor The setting up of the EMR was based on our experiences of its application to enzymatic hydrolysis of vegetable oils and fats. The membrane tubes were fixed by epoxy glue and it has been proved to bc fairly resistant to the chemicals applied [5]. Unfortunately, it was found during the esterification runs that the ethyl oleate/oleic acid mixture causes the corrosion of the glue. Due this effect, the aqueous and organic phases were mixed after 100-200 hours and the experiments had to be stopped. After testing the possibilities to solve this problem, a more advanced reactor structure with flat membrane has been elaborated for the further investigations. 4. C O N C L U S I O N S
Based on the analysis of these studies,conclusions can be drawn as follows: Comparing the differentimmobilization methods of lipasc (Table I) significantadvantage can bc found for the coating and emulsion procedures. Although the same method was used in the last two cases, the difference can probably be explained by the considerable different medium composition and conditions. A relativelyhigh ester concentration can be achieved in the secondary side of the enzyrnc membrane reactor even in case of low (30-40) ethanol concentrations in the primary side. If the operational stabilityof E M R will bc improved by the application of the reactor of new type, presumably flavour esteryields can be enhanced further. The presence of living cells and other components of the broth did not reduced dramatically the ester formation rate. Consequently, the further studies on extractive bioconversion reactor system consisting of the immobilized cell fermenter and E M R seem to bc promising for natural flavour esterproduction.
106 Table 1. Values of initial ester formation rate related to membrane surface area
Medium in the primary side
Immobilization method
Ethyl oleate formation rate (g day"m"2).........
Aqueous ethanol
UF/adsorption
239
Aqueous ethanol/ glucose/biomass
Coating
638
Aqueous ethanol/ glucose/biomass
Emulsion
854
Fermentation broth with living cells
Emulsion
414
REFERENCES
1. 2. 3. 4. 5.
D.R. Wu, S.M. Cramer and G. Belfort, Biotechnol. Bioeng. 41 (1993) 979. K. B61afi-Bak6, A. Dombi, L. Szab6 and E. Nagy, Biotechnol. Techniques. 8 (1994) 671. S.T. Bouwer, F.P. Cuperus and J.T.P. Derksen, Enzyme Microb. Tehnol. 21 (1997) 291. M. Habulin and Z. Knez, J. Membr. Sci. 61 (1991) 315. E. Cshnyi, K. B61afi-Bak6, T. Venyige and Cs.Sisak, Proc 1 st E u r . Congr. on Chem Eng. Florence. Vol 4. (1997) 2627. 6. M.R. Aires-Barros, J.M.S. Cabral and J.M Novais, Biotechnol. Bioeng. 29 (1987) 1097. 7. A.C. Oliveira, and J.M.S. Cabral, J. Chem Tech. Biotechnol. 52 (1991) 219.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
107
Study of the deactivation process of the glucose oxidase-catalase enzymatic system by m e a n s of simulation L. E. Romero* and D. Cantero Biological and Enzymatic Reactors Research Group. Department of Chemical Engineering, Food Technology and Environmental Technologies. Faculty of Sciences. University of Cadiz. 11510. Puerto Real. Cadiz. Spain.
This paper reports the simultaneous deactivation of co-immobilized glucose oxidase and catalase in calcium alginate beads. For this purpose, the study is conducted in a continuous stirred tank reactor containing the reaction medium. After a comprehensive review of the existing literature, a three-parametered irreversible deactivation general model is put forward for each enzyme. 1. INTRODUCTION Nowadays, glucose oxidase (GOD) and catalase (CAT) are considered to be one of the enzymatic systems of most interest and use for both sensors and industrial processes. Glucose oxidase and catalase are nature related because they both participate in the enzymatic pool of those microorganisms able to oxidize glucose to gluconic acid [1]. Glucose oxidase has the systematic name of 13-D-glucose: oxygen-l-oxidoreductase (EC 1.1.3.4.) and catalase is H202: 1-1202 oxidoreductase (EC 1.11.1.6.). Both enzymes (GOD) and (CAT) are very stable against pH and temperature; consequently, their pH range is very wide (pH 3-8) [2,3]. If temperature of operation is considered, both enzymes show catalytic activity from 10~ to 60~ [2, 3, 4]; however, they are highly sensitive against hydrogen peroxide [5]. GLUCOSE~
/ [
3N
+
+
DEACTIVATIO~ HYDROGEN- l
Figure 1. Hydrogen peroxide deactivating effects.
]
108 1.1. Deactivation mechanism of glucose oxidase. Existing literature provides evidence for a "ping-pong" model to be used in the irreversible deactivation of the GOD enzyme [5]. Such model assumes that hydrogen peroxide works by deactivating the three states this enzyme is likely to adopt: oxidized state, reduced state, and complex GOD-oxygen state. Further to enzyme activity and reaction time, a representative equation would be:
a=exp(-klP 0
(1)
where: a: relative activity of the enzyme (between 0 and 1) k1: deactivation coefficient. P: concentration of hydrogen peroxide. t: time of reaction. 1.2. Deactivation mechanisms of catalase.
Two different models have been employed for the deactivation mechanisms of catalase, i.e. Tse and Gough's [5], and Vasudevan and Weiland's [6]. Tse and Gough's model is similar to that for GOD. This means that the enzyme is deactivated in the oxidized state, in the reduced state, as well as in the complex enzyme-peroxide state. It provides the following equation: a = exp
(-kot - kiP t)
(2)
In this equation, a new kinetic constant appears, this is k~ Such constant is not affected by hydrogen peroxide concentration. Finally, the model by Vasudevan and Weiland [6] assumes that there exist three states of the enzyme. These states are: enzyme, enzyme-peroxide, and enzyme-peroxide-peroxide. A mathematical treatment of this mechanism provides the following equation:
a = exp
-
klpn p2 t
(3)
P+k 2 +~ k3 Where kl, k2 and k~ are deactivation constants. It can be observed that, whereas the mathematical expression is more complicated than the previous one, the mathematical function is very similar. 2. MATERIALS AND METHODS 2.1. Reactants Experiments were developed by the use of glucose oxidase from SIGMA (G-1262, crude extract). The enzyme had 10.600 A.U./g solid for conditions of oxygen saturation (pH 5.1 and 35~ Catalase employed was from SIGMA (C-3515) with 7080 A.U./mg solid.
109 2.2. Reaction medium The experiments were developed using D-(+)-glucose from PANREAC dissolved in a pH 5.1 buffer (acetic acid/calcium acetate). 2.3. Calcium aiginate gel (CAG) beads C AG beads were prepared by means of the reaction of alginic acid and calcium chloride. A general technique of ionotropic gelling was used for this purpose [7]. Calcium alginate gel beads behaved as carriers for both glucose oxidase and catalase. 2.4. Reactor and starting up The reactor used was a 300 mL stirred tank reactor with a working volume of 250 mL. All the experiments were developed in continuous operation, working at different residence times and with several substrate initial concentrations. The temperature of operation was 35~ and pH was 5.1 -as these are regarded as the optimum operating conditions for glucose oxidase. 2.5. Software The simulation software was developed in C-language using LabWindows (2.2.1.) from National Instruments, and it was finally compiled by Microsoft (6.0). The developed software acquires process variables data and parameters related with the calculus algorithm (finite differences method), providing the simulation results. These results can be simultaneously filed and displayed [8]
' I-'i o
REACTION RATES l
~ J CAT P"I
MASS BALANCES EFFECTIVENESS FACTORS SUSTRATE
PRODUCT
1
! ~176
l l|l
ii
i
I
Figure 2. KINETIC M O D E L FOR THE. SYSTEM
3. RESULTS AND DISCUSSION. As a result of in-depth experimental studies concerned with the kinetic behaviour of GOD and CAT immobilized on CAG beads, a kinetic model is finally proposed. This model is made up of several equations, such as (i) equations for the enzymes (reaction rates and deactivation); (ii) mass balances for substrate and product both inside the particles and inside the reactor; (iii)
110 conversion and effectiveness factors. Unlike traditional models applied to enzymatic processes [9], the model put forward here shows deactivation equations. After a comprehensive review of the existing literature, a three-parameters irreversible deactivation general model is put forward for each enzyme:
aGO D = e x p -
kI(GoD)P2
t , aCA T exp " = -
t
k3(GOD ) + P
k3(cAr) + P
Where: "' ac,oD" and "'acAr " are the relative activities of GOD and CAT; "k" 1 to 3 are the deactivation coefficients for each enzyme; "P" is the hydrogen peroxide concentration; and "t" is the time of reaction. The adjusted values of these coefficients are: kl(c, oD) = 5- 10 "5 (mM s) "I kI(CAT) = 3- 10"2 (mM s) "1 k2(GOD) =
2. 10 "3 s"1
k2(CAT)'-5- I0 3 s~
ka(c, oD) =
1.0 mM
k3tCAT)= 1.2 mM
With the aim of obtaining the adjusted values of deactivation coefficients, these two deactivation equations were included iy: a simulation program together with all those equations necessary to predict the behaviour of tl '~. system. Once developed, it was possible to validate the simulation program thanks to a vast experimentation carried out. Results obtained facilitate the knowledge of the enzymatic oxidation process from glucose to gluconic acid and hydrogen peroxide, starting from its initial stage along the whole process until the complete deactivation of the system.
A
E 15 r
Z
o .,..,
@
9
I--
~ 10 I-, Z
o z o o
5
0
6
12
18-
24
30
36
42
TIME (h)
Figure 3. Experimental substrate concentration ( t ) , experimental product concentration (11); theoretical substrate concentration ( ) and theoretical product concentration ( . . . . ) versus time.
111 In Figure 3 it is possible to check the experimental and simulated concentrations of glucose (susbstrate) and hydrogen preroxide (product) in the reaction medium. As it can be observed, the system works at peak performance along the first ten hours of the process. Then, the system gradually begins to deactivate, substrate concentration returns to its inital value, and product concentration finally falls down to zero due to the wash-out effect. The developed simulation software allows for the calculation of the deactivation profiles of both enzymes GOD and CAT (Figures 4 and 5). Thus, it can be observed that relative activities of the enzymes can be predicted with the same equation, although their behaviour is completely different for each case. Figure 4 shows simulated profiles of relative activity versus dimensionless radius for different times. It should be observed that at the surface of particles r = 0 and at the center of particles r = 1. The enzyme glucose oxidase follows a "progressive conversion" model in which activity decays with the same degree in all radial positions. This behaviour is explained by an irreversible enzyme deactivation which increases along with the process time. Thus, the time the enzyme is located in an environment with unfavourable conditions seems to be the most relevant factor in the deactivation kinetics.
,
2h 0.8
Q I
r=l
)0.6
(3
Oh
r=O
5h 0.4
0.2 .I 0
25h
1Oh ,
,
0.2
0.4
'
! 0.6
0.8
DIMENSIONLESS RADIUS (r)
1
Figure 4. Simulated profiles of relative activity GOD versus dimensionless radius.
On the other hand, catalase deactivation (Figure 5) can be represented by a "decreasing nucleus" model from the surface to the center of the particles. Figures 4 and 5 also contribute to conclude that CAT deactivates earlier than GOD. This way, the relative activity of catalase is zero after five hours of process and, at this point, GOD still maintains a 60% of its activity (relative activity 0.6). As a conclusion, a sequential deactivation of both co-immobilized enzymes can be observed: CAT decays in the first place, and GOD in the second place, each one with a different deactivation model.
112
0.8
0
0
3h
I-" 0.6
4h
0.4 0.2
0
0.2
0.4
0.6
0.8
1
DIMENSIONLESS RADIUS (r) Figure 5. Simulated profiles of relative activity CAT versus dimensionless radius.
REFERENCES 1. R6hr, M., Kubicek, C.P. and Kominek, J. Biotechnology 0Ed. Rehm, H.J. and Reed, G.) Verlag Chemic Gmbh, Weinheim (1983). 2. Kozhukharova, A., Kirova, N., Popova, I. Batsalova, K. and Kunchev, K.. Biotechnol. Bioeng,, 32 (1988) 245. 3. Jiang, B. and Zhang, I.. Eur. Polym. J., 29 (1993) 1251. 4. Fortier, G. and Bdanger, D. Biotechnol. Bioeng., 37 (1991) 854. 5. Tse, H.S.P. and Gough, D.A. Biotechnol. Bioeng., 29 (1987) 705. 6. Vasudevan, P.T. and Weiland, R.H. Biotechnol. Bioeng,, 36 (1990) 783. 7. Smidsr~d, O. and Skjg~k-Bra~k,G. TmTECH, 8 (1990) 71. 8. Romero, L.E.. PhD. Thesis. University of Cadiz, Spain (1996). 9. B6dalo, A., G6mez, J.L., G6mez, E., Bastida, J. and Martinez, E. Biotechnol. Prop=, 9 (1993) 166.
Stabilization by chemical modification
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
115
Stabilization of hydrolases by chemical modification with fatty acids or polyethylene glycol F.J. Plou, M.V. Calvo, M. F e r r e r a n d A. B a l l e s t e r o s
Departamento de Biocat~isis, Instituto de Cat~lisis, CSIC, Campus Universidad Aut6noma, 28049 Madrid, Spain 1. I N T R O D U C T I O N One of the m o s t i m p o r t a n t aims of enzyme engineering is to e n h a n c e the stability of enzymes. In particular, the t e m p e r a t u r e stability of e n z y m e s u s e d in l a u n d r y d e t e r g e n t s (lipases, proteases, amylases) is a n i m p o r t a n t factor in productivity. Protein stabilization h a s b e e n achieved by several m e t h o d s (Shami et al., 1989), including immobilization, site-directed m u t a g e n e s i s , chemical modification, crosslinking a n d selection of e n z y m e s from thermophilic organisms. Chemical modification of an enzyme m a y lead to a n alteration of its surface and can t h u s be u s e d to i m p a r t d e s i r e d properties to the biocatalyst. The hydrophile-lipophile b a l a n c e (HLB) of a n enzyme s e e m s to be i m p o r t a n t in some of its properties (recognition of s u b s t r a t e s , binding, stability, etc.). In this context, the in vivo p o s t - t r a n s l a t i o n a l modification of proteins by the covalent a t t a c h m e n t of fatty acids a n d lipids is a w i d e s p r e a d p h e n o m e n o n in n a t u r e b u t as yet not completely u n d e r s t o o d (Schultz et al., 1988). Several m e t h o d s for the acylation of proteins in vitro have b e e n also described. Polyethylene glycol (PEG) is a synthetic p o l y m e r with a m p h i p a t h i c properties. The hydrophilic n a t u r e of P E G m a k e s it possible to modify e n z y m e s in a q u e o u s solution, a n d its h y d r o p h o b i c c h a r a c t e r e n a b l e s the modified e n z y m e s to function in h y d r o p h o b i c e n v i r o n m e n t s . PEG-modified e n z y m e s have b e e n reported to be soluble a n d active in a p o l a r organic solvents. In c o n t r a s t with the v a s t n u m b e r of studies on synthetic or t h e r a p e u t i c a l applications of PEG-enzymes, the effect of this modification on enzyme stability h a s b e e n analyzed by only a few r e s e a r c h e r s {Pasta et al., 1988; Calvo et al., 1995}. Subtilisin is a serine e n d o p e p t i d a s e (MW 27500) t h a t is secreted in large a m o u n t s from a wide variety of Bacillus species. The large d a t a b a s e for subtilisin m a k e s it a n attractive model enzyme to elucidate s t r u c t u r e - f u n c t i o n relationships. The lipase p r o d u c e d by Candida rugosa (formerly C andida cylindracea ) is being extensively u s e d in research, owing to its high activity in hydrolytic as well as in synthetic reactions (Otero et al., 1990). This y e a s t p r o d u c e s two m a i n isoforms (lipases A a n d B) with similar a m i n o acid content, N-terminal s e q u e n c e a n d molecular weight, w h i c h differ in n e u t r a l s u g a r content, hydrophobicity, stability We thank Comunidad de Madrid and Fundaci6n Caja de Madrid for financial support. This research was financed by the Spanish CICYT (project BIO96-1574-CE).
116 and substrate specificity (R~a et al., 1993). Our group h a s developed a novel procedure for the purification of lipases A and B based on hydrophobic interaction chromatography (Rua and Ballesteros, 1994). In the present work, and imitating the hydrophobization that takes place in n a t u r e for m a n y proteins, we have lowered the HLB of subtilisin Carlsberg - a protein containing 9 L y s + the amino terminal (Ala)- by chemical acylation using acyl chlorides of different chain length. In addition, we have modified lipases A and B from C. rugosa by covalent coupling of their amino groups with PEG activated by two different m e t h o d s (see Fig. 1). The effect of chemical modification on the activity and stability of the corresponding enzymes was further analyzed.
CH'(OCH'CH2'"O c' O--NH,
CH3 (OCH2CH2)nO~_ .... N
CH3 (OCH2CH2)n O ' ~ ~ N
OH3 (OOH2OH2)nO ~
O
CHa(OCH2CH2)nO~O
(~NH
~
2
+ HCI (I)
0
~~'--NH~)
CHz(OCI'I2CH2)nO~;--NH-~
+ HO--
([B
O-~
o CH3 (CH2)n CII--Cl
o ~--- NH2 --- CH3 (CH2)nCII--NH~[~
+ HCI (IH)
Figure 1. Scheme of the chemical modification of enzymes with: (I) methoxypolyethylene glycol activated with triazine (TZ-PEG); (II) methoxy-polyethylene glycol activated with succirdmidyl carbonate (SC-PEG); (III) fatty-acid chlorides.
2. MATERIALS AND METHODS Subtilisin Carlsberg (protease type VIII from B. licheniformis}, palmitoyl and octanoyl chlorides, sodium cholate, p-nitrophenyl butyrate (pNPB), tributyrin, Succ-Ala-Ala-Pro-Phe-pNA, methoxy-polyethylene glycol (MW 5000) activated with cyanuric chloride (TZ-PEG) a n d C. rugosa lipase were from Sigma. Methoxypolyethylene glycol (MW 2000 and 5000) activated with succinimidyl carbonate (SC-PEG) was from Calbiochem. 2,4,6-Trinitro-benzenesulfonic acid and PMSF were from Fluka. Phenyl-Sepharose and S e p h a d e x G - 5 0 were supplied by Pharmacia. Sodium dodecyl sulfate (SDS) was from Serva.
2. I. Lipase p u r i f i c a t i o n Lipase from C. rugosa was purified using an FPLC system (Pharmacia) into isoenzymes A and B following the protocol of Rua and Ballesteros (1994) with Phenyl-Sepharose as matrix.
117
2.2. E n z y m e a s s a y s The a m i d a s e activity of subtilisin was d e t e r m i n e d spectrophotometrically at 410 n m in 0.1 M phosphate buffer (pH 7.0), containing 0.1 M Succ-Ala-Ala-Pro-Phe-pNA a s s u b s t r a t e . The hydrolytic activity u s i n g p-nitrophenyl b u t y r a t e a s s u b s t r a t e was followed spectrophotometricaUy at 346 nm. The a s s a y mixture contained 0.32 mM s u b s t r a t e in 50 mM p h o s p h a t e buffer (pH 7.2), including 4% acetone. Lipase activity u s i n g tributyrin was a s s a y e d titrimetrically in a pH-stat {Radiometer) u s i n g 5 0 r a M NaOH. The reaction mixture contained 68 mM tributyrin, 0.1 M NaC1, 0.1 M CaC12, i mM Tris-HC1 (pH 7.2), including 3% acetonitrile. One u n i t of activity (U) is the a m o u n t of enzyme t h a t hydrolyses 1 pmol of s u b s t r a t e / m i n u n d e r the above conditions.
2.3. C h e m i c a l m o d i f i c a t i o n The acylation of subtilisin w a s performed in a p H - s t a t according to a method previously d e s c r i b e d (Plou a n d Ballesteros, 1994). Palmitoyl chloride and octanoyl chloride were employed as acylating agents; the former was u s e d in p r e s e n c e of sodium cholate to p r o d u c e a n h o m o g e n e o u s dispersion. Chemical modification of lipases with activated PEG was carried out using a pH-stat. Lipase (25 mg) w a s dissolved in 6 mL of 50 mM p h o s p h a t e buffer {pH 8.0). Then, PEG w a s a d d e d a n d the mixture m a i n t a i n e d at pH 8.0 and 25~ with 50 mM NaOH. The m o l a r excess of activated PEG referred to the amino g r o u p s in the protein was 10:1. Once the b a s e c o n s u m p t i o n curve leveled off, the u n b o u n d PEG was removed by ultrafiltration - u s i n g an Amicon PM 30 m e m b r a n e - w a s h i n g with 1 mM p h o s p h a t e buffer (pH 7.0). Finally, the resulting PEG-lipase was freeze-dried.
2.4. D e t e r m i n a t i o n of t h e d e g r e e of s u b s t i t u t i o n The c o n t e n t of free lysines before a n d after acylation with fatty acids was determined u s i n g 2,4,6-trinitrobenzenesulfonic acid according to Fields (1971}. In order to avoid the effect of the p e p t i d e s g e n e r a t e d by autolysis of subtilisin, leading to an o v e r e s t i m a t i o n of a m i n o groups, s a m p l e s were inhibited irreversibly with PMSF and c h r o m a t o g r a p h i e d over S e p h a d e x G-50. The c o n t e n t of a m i n o g r o u p s in the protein before and after the chemical modification with PEG w a s d e t e r m i n e d by the fluorescamine m e t h o d , using a Perkin Elmer LS50B spectrofluorometer, as described by Stocks e t al. (1986). F l u o r e s c a m i n e was b o u n d to the free a m i n o groups, and the intensity of the fluorescence e m i s s i o n was m e a s u r e d at 475 n m after excitation at 390 nm.
2.5. A s s a y o f s t a b i l i t y A diluted solution (1-2 ~tM) of enzyme in the corresponding buffer {in absence or p r e s e n c e of SOS) w a s i n c u b a t e d at the desired t e m p e r a t u r e . The r e m a i n i n g activity with Succ-Ala-Ala-Pro-Phe-pNA for subtilisin or with pNPB for C. r u g o s a lipase was m e a s u r e d at intervals. The c o r r e s p o n d i n g curves were fitted to a single exponential (subtilisin, lipase B} or a double exponential decay (lipase A) using the SIMFIT p a c k a g e {Bardsley, 1995}. Buffers employed were 0.1 M p h o s p h a t e (pH 7.0) for subtilisin, a n d 50 mM p h o s p h a t e (pH 7.2) or 0.1 M b i c a r b o n a t e (pH 9.0) for lipases.
118
3. RESULTS AND DISCUSION 3.1. Acylation of subtilisia with fatty acids The kinetic c o n s t a n t s for the hydrolysis of n a t u r a l a n d synthetic s u b s t r a t e s catalyzed by native a n d modified subtilisins were evaluated. Slight differences in kcat a n d Km were o b s e r v e d in the hydrolysis of synthetic s u b s t r a t e s . However, u s i n g n a t u r a l s u b s t r a t e s - s u c h as c a s e i n - the chemical modification p r o d u c e d a d e c r e a s e in the catalytic efficiency, possibly d u e to steric h i n d r a n c e ; the effect w a s more appreciable with palmitoyl-subtflisin c o m p a r e d with octanoyl-subtilisin (Plou a n d Ballesteros, 1994) The stability of acyl-subtilisins in a q u e o u s m e d i u m (pH 7.0) at 45~ a n d 65~ w a s studied. Low enzymatic c o n c e n t r a t i o n s were u s e d in order to minimize autolysis a n d / o r aggregation processes. As s h o w n in Table 1, the h y d r o p h o b i z a t i o n of subtilisin with acyl chlorides (C8 or C16) e n h a n c e s significantly its stability. The stabilization effect (ratio of the t h e r m o i n a c t i v a t i o n c o n s t a n t s of native a n d modified enzymes) is a b o u t 1S-fold at 45~ a n d a b o u t 2.S-fold at 65~ Table 1 Half-lives (t112) of native a n d fatty-acid modified subtilisins at pH.7 Fatty acid
Degree of
tl/2 (h)
acylationl (%)
45~
65~
Native
--
2.4
1.4
Octanoic acid
57
30.4
2.9
Palmitic acid
60
37.2
4.4
1 Subtilisin Carlsberg c o n t a i n s 10 a m i n o residues.
3 . 2 . M o d i f i c a t i o n o f l i p a s e s w i t h PEG Chemical modification of the a m i n o g r o u p s of lipases A a n d B with PEGtriazine (TZ-PEG) a n d PEG-succinimidyl c a r b o n a t e (SC-PEG) w a s carried out. S u b s t i t u t i o n degrees r a n g i n g from 5 0 - 7 0 % were achieved (lipases A a n d B c o n t a i n 25 a n d 21 lysine r e s i d u e s per mol respectively). It is i m p o r t a n t to note t h a t w h e n PEGsooo is activated with triazine, two c h a i n s of PEG are b o u n d to the biocatalyst per single s u b s t i t u t i o n {Fig. 1). The stability of TZ-PEGsooo-lipases A a n d B w a s studied at different v a l u e s of pH. At pH 7.2, the modification does not p r o d u c e a significant stabilization {results not shown). However, at pH 9.0, the TZ-PEGsooo-lipases A a n d B are c o n s i d e r a b l y more stable t h a n the native enzymes. Fig. 2 s h o w s the inactivation curves for the native a n d modified l i p a s e s A a n d B, respectively. The stabilizing effect, in t e r m s of the inactivation c o n s t a n t s , is 9-fold for lipase A a n d 20-fold for lipase B. Similar stabilization effects were f o u n d u p o n modification with SC-PEG. The stability of native a n d chemically-modified p r e p a r a t i o n s of lipases A a n d B i n c u b a t e d with v a r i o u s c o n c e n t r a t i o n s of SDS (0.04-0.5%) w a s also examined. For lipase A, the modification with PEG gives rise to p r e p a r a t i o n s with higher stability
119 (about 4-fold); however, the m e t h o d of activation of PEG w a s less i m p o r t a n t t h a n the degree of s u b s t i t u t i o n in this effect. 100 ~"
,--
80
~
,-,9
o
60
4O =1 .,.-i
m
4 Z L ~
20 0 1
2
3
4
5
6
T i m e (h) Figure 2. Kinetics of e n z y m e inactivation at pH 9.0 o b s e r v e d with lipase A (O), TZPEOsooo-lipase A ([:]), lipase B (O) a n d a n d TZ-PEGsooo-hpase B (tt). The i n a c t i v a t i o n of l i p a s e B at 0 . 2 % SDS w a s so fast t h a t it w a s not possible to d e t e r m i n e a n a c c u r a t e v a l u e for kin. Fig. 3 s h o w s the inactivation c u r v e s for lipase B a n d PEG-lipase B at 0 . 2 % (w/v) SDS, r e p r e s e n t e d in a s e m i l o g a r i t h m i c scale. Fig. 4 s h o w s the i n a c t i v a t i o n c u r v e s for lipase B a n d PEG-lipase B at 0 . 0 4 % (w/v) SDS. At both c o n c e n t r a t i o n s of SDS, the l i p a s e s modified with PEGsuccinimidyl c a r b o n a t e are m o r e stable t h a n t h o s e with PEG-triazine, especially at 0.04% (w/v) SDS. In all c a s e s , the modification with PEG stabilizes significantly lipase B with r e g a r d s to the d e l e t e r i o u s effect of SDS. The b i p h a s i c c h a r a c t e r of the inactivation c u r v e s of s a m p l e s u s i n g SC-PEG m i g h t reflect the p r e s e n c e of p o p u l a t i o n s of PEG-lipase with different d e g r e e s of s u b s t i t u t i o n . 100
> ~
10
a)
x 0
i 25
I
1
1
50
75
I00
Time
125
(min)
Figure 3. S e m i l o g a r i t h m i c plot of the inactivation of lipase B (O} a n d the s a m p l e s modified TZ-PEGsooo (A), SC-PEGsooo (O) a n d SC-PEG2ooo (0) i n c u b a t e d in a q u e o u s solution at 25~ a n d pH 7.2, in p r e s e n c e of 0.2 % (w/v) SDS.
120 1 O0
L.
~
~
.~
I
~
~
i
80 .+a .P,I
.~9 .l.a {j
60 4O
"'~ ~)
ID
20
...
0
1
2
I
3
4
Time
5
6
7
8
(h)
Figure 4. Stability of lipase B (O) and the samples modified TZ-PEGsooo (A), SCPEGsooo (O) and SC-PEG2ooo (0) incubated in aqueous solution at 25~ and pH 7.2, in presence of 0.04 %(w/v) SDS. In conclusion, chemical modification of enzymes with fatty acids or amphipatic polymers such as PEG may be exploited as a means of obtaining preparations with enhanced stability u n d e r harsh conditions: high temperature, alkaline pH and presence of anionic surfactants. The increased resistance of acyl-subtilisin and PEG-lipases to these d e n a t u r a n t s is of great interest from the point of view of structural implications a n d / o r the biotechnological potential of hydrolytic enzymes.
REFERENCES
Bardsley, G.W. (1995) SIMFIT: "A package for simulation, curve-fitting and statistical analysis in the life sciences", Manchester University, U.K. Cairo, M.V., Plou, F.J., Pastor, E. and BaUesteros, A. Biotechnol. Lett. 17 (1995) 171-176. Calvo, M.V., Plou, F.J. and BaUesteros, A. Biocatal. Biotransf. 13 (1996) 271-285. Fields, R. Biochem. J., 124 (1971) 581-590. Otero, C., Pastor, E., Fernandez, V.M. and Ballesteros, A. Appl. Biochem. Biotechnol. 23 (1990) 237-247. Pasta, P., Riva, S. and Carrea, G. FEBS Lett. 236 (1988) 329-332. Plou, F.J. and BaUesteros, A. FEBS Lett. 339 (1994) 200-204. Rfla, M.L., Diaz-Maurifio, T., Fem~_ndez, V.M., Otero, C. and Ballesteros, A. Biochim. Biophys. Acta 1156 (1993) 181-189. Rfia, M.L. and BaUesteros, A. Biotechnol. Tech. 8 (1994) 21-26. Schultz, A.M., Henderson, L.E. and Oroszlan, S. Annu. Rev. Cell Biol. 4 (1988) 611-647. Shami, E.Y., Rothstein, A. and Ramjeesingh, M. Trends Biotechnol. 7 (1989) 186-190. Stocks, S.J., Jones, A.J.M., Ramey, C. W. and Brooks, D.E. Anal. Biochem. 154 (1986) 232-234.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborraand P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
121
M o d i f i c a t i o n o f t h e c a r b o h y d r a t e m o i e t y in r i b o n u c l e a s e B a n d its influence on the protein stability probed by limited proteolysis
Ulrich Arnold and Renate Ulbrich-Hofmann Martin-Luther University Halle-Wittenberg, Department of Biochemistry and Biotechnology, Institute of Biotechnology, Kurt-Mothes-Strat~e 3, D-06120 Halle, Germany
The non-glycosylated ribonuclease A (RNase A), its glycosylated form, ribonuclease B (RNase B), and two partially deglycosylated species of RNase B were compared with respect to their kinetic thermal stabilities determined by limited proteolysis with thermolysin. The modifications of the carbohydrate moiety in ribonuclease B were obtained by treatment with endoglycosidase F and c~-mannosidase, respectively, yielding GlcNAc-RNase and GlcNAc2Mart3-RNase (Man3-RNase). The results reveal that the modified RNase species possess nearly the same stability as RNase B independent of the size of the carbohydrate moiety. In contrast, the rate of proteolysis decreased with increasing volume of the carbohydrate chain due to steric shielding of the primary cleavage sites in the ribonuclease molecule. 1. INTRODUCTION Glycosylation as the mostly occurring natural modification of proteins is suggested to function as traffic marker, signal modifier, or regulator of biological activities (Rademacher et al., 1988; Lis & Sharon, 1993). Moreover, it is known to effect the stability of the protein structure toward denaturants (Jaenicke, 1991) and proteolytic degradation (Rudd et al., 1995). Glycosylation might increase the life time of the protein molecule by prevention of deamidation of Asn side chains which results in a decreased stability of the protein (Catanzano et al., 1997). Ribonuclease A (RNase A) and ribonuclease B (RNase B) possessing identical primary and tertiary protein structures (Berman et al., 1981; Williams et al., 1987) but differing by a carbohydrate chain (GlcNAc2Mans_9) attached to Asn34 in RNase B are an ideal model system for studying effects due to glycosylation. Interestingly, the glycosylation site is situated in the structural region that was identified as the first unfolding region in RNase A under thermal denaturation (Arnold et al., 1996). The location of the unfolding region is not changed due to the attachment of the carbohydrate moiety in RNase B
122 (Arnold et al., 1998) whereas the kinetic as well as thermodynamic thermal stabilities of RNase B are higher than those of RNase A (Arnold & Ulbrich-Hofmann, 1997). In the present paper, the influence of the size of the carbohydrate moiety on the kinetic thermal stability and the proteolytic resistance of the RNase molecule is studied. Therefore, partially deglycosylated RNase B species were prepared and their unfolding and proteolyis rate constants were determined by limited proteolysis with thermolysin.
o Ash34 RNase A
0 GlcNAc
'~ Man
GlcNAcRNase
Man3RNase
RNase B
Figure 1. Schematic presentation of the modification of the carbohydrate moiety in the RNase molecule. 2. MATERIALS A N D METHODS
RNase A and RNase B from Sigma, MO, USA, were purified on a MONO S FPLCcolumn (Pharmacia Biotech, Sweden) using 50mM Tris-HC1 buffer, pH8.0. c~-mannosidase and thermolysin from Sigma, MO, USA, and endoglycosidase F from Boehringer, Germany, were used without further purification. All other chemicals were the purest ones commercially available. Preparation of GlcNAc-RNase and Man3-RNase. After adjusting the pH to 4.5 with 1 M sodium acetate buffer, 1 mg of purified RNase B were treated with endoglycosidase F (0.5 U) or c,-mannosidase (1.0 U) at 35~ for 22 h. The reaction products were purified on a MONO S FPLC-column and identified by MALDI mass spectrometry (Reflex TM Bruker-Franzen, Germany). Thermal denaturation and proteolysis. In a typical experiment, 15 ~tL of thermolysin (0.1-0.8 mg/mL) in 50 mM Tris-HC1 buffer, 10 mM CaC12, pH 8.0, and 15 I~L of RNase (1.0 mg/mL) were added to 120 ~tL of preincubated 50 mM Tris-HC1 buffer, pH 8.0. After distinct time intervals, samples of 18 I~L were removed and mixed with 6 btL of 50 mM EDTA.
123 Electrophoresis and densitometric evaluation. Electrophoresis was carried out on a Midget electrophoresis unit (Hoefer) according to Sch~igger & v o n Jagow (1987) but using 10, 14, and 18% acrylamide for the sampling, spacer, and separating gels. The SDS-PAGE gels were stained with silver nitrate according to Blum et al. (1987) or stained with Coomassie Brillant Blue G250 and scanned at 595 nm using a densitometer CD 60 (Desaga). Determination of the unfolding rate constants. The rate constants of proteolysis (kproteolysis) w e r e calculated for several thermolysin concentrations from the decrease of the peak areas of intact RNases in the scanned SDS-PAGE gels as a function of time of proteolysis according to Arnold and Ulbrich-Hofmann (1997). From the respective pairs of kproteolysis and the thermolysin concentration [P]0, the unfolding rate constants (ku) were estimated in adoption to the approach by Imoto et al. (1986) with
ku [P]0 kproteolysis=
and B + [P]0
lim kproteolysis= ku [Pl0-*~
where B is a constant. 3. RESULTS The carbohydrate moiety of RNase B was shortened by its treatment with endoglycosidae F or ~-mannosidase, which are known to cleave the oligosaccharide chain between the di-N-acetylchitobiose moiety and at Man~l--~X bonds, respectively. The partial deglycosylation and purification procedures of the ribonucleases led to homogenous products in the SDS-PAGE (Fig. 2) which were identified as GlcNAc-RNase and Man3-RNase by MALDI mass spectrometry. Surprisingly, the deglycosylation of RNase B by c~-mannosidase did not lead to Manl-RNase but reaction stopped at the Man3 level. This result is in contrast to the results by Rudd et al. (1994) where, however, different reaction conditions were applied.
Figure 2. SDS-PAGE of RNase B (lane 1), Man3-RNase (lane 2), GlcNAc-RNase (lane 3), and RNase A (lane 4). Lane 5 shows the reference proteins soybean trypsin inhibitor (21 kDa), cytochrom c (12.4 kDa), and bovine pancreatic trypsin inhibitor (6.5 kDa).
124 For comparing the kinetic thermal stabilities of the different glycosylated RNase species, the method described in Arnold and Ulbrich-Hofmann (1997) was applied. Both the naturally occurring RNases and the partially deglycosylated RNases were treated with thermolysin of different concentrations under thermal denaturing conditions (50-60~ In all cases, increasing thermolysin concentrations as well as increasing temperatures led to increased proteolysis rates for the RNases. Even if the proteolytic degradation of the glycosylated RNase species was markedly decelerated by the more or less voluminuous carbohydrate moieties, the rate constants of proteolysis revealed a saturation behavior with increasing thermolysin concentrations. Therefore, the unfolding rate constants could be estimated by extrapolation of the proteolysis rates to infinite protease concentrations (Table 1). Table 1.
Unfolding rate constants (ku) of modified and non-modified RNases (xl0 -4 s-l). T (~ 50 55 60
RNase A
GlcNAc-RNase
Man3-Rnase
RNase B
8.25 + 0.95 39.6 + 2.8 159 + 21
2.39 + 0.17 18.6 + 1.3 92.7 + 10
2.74 + 0.19 21.4 +1.8 77.6 + 5.8
2.66 + 0.17 17.5 + 5.2 76.5 + 10.9
Figure 3. EYRING-plot of the unfolding rate constants of the modified and non-modified RNases ( 9 RNase A, 9GlcNAcRNase, ~ Man3-RNase, and 9RNase B).
Within the experimental error, all the non-RNase A species reveal the same unfolding rate constants, which clearly differ from those of RNase A at all temperatures studied. From the EYRING-plot of the ku-values (Fig. 3) the activation enthalpy AH# and the activation entropy AS# were estimated to be 263 + 5 kcal moN
125 and 508 + 9 cal mo1-1 K -1 for RNase A and 298 + 7 kcal mo1-1 and 608 + 23 cal mo1-1 K -~ for RNase B. Nearly identical values to those of RNase B were obtained for both the modified species. On the other hand, the proteolytic susceptibility of the different glycosylated RNase species was compared under conditions where the concentration of thermolysin and not the rate of unfolding of the RNases limits the rate of proteolytic degradation, i.e. at high temperatures and low protease concentration (Fig. 4). The results demonstrate that the rate of proteolysis is strongly influenced by the modification of the ribonuclease molecule. 100 -
Figure4. Time course of the proteolytic degradation of ribonucleases by thermolysin at a 250:1 ratio ( w / w ) at 65~ The kproteolysis -values are 8.46 x 10 -3 s -1, 7.23 x 10 -3 s-1, 5.13 x 10 .3 s -1, and 4.33 x 10 -3 s-1 for R N a s e A ( - ) , GlcNAcRNase ( 9), Man3-RNase ( a ), and RNase B ( 9).
50
t~- "
10
._.
\
r
~ ..Q 0 (1,}
2
0.5
l_
0.1
0
i
i
i
250
500
750
1000
time (s)
4. D I S C U S S I O N Limited proteolysis proved to be a powerful tool both for the detection of delicate changes of the protein structure due to modifications and the determination of the kinetic stability. The comparison of the four RNase species revealed that the proteolytic degradation which starts at the Asn34-Leu35 and the Thr45-Phe46 peptide bonds (Arnold et al., 1996,1998) is hindered by the glycosylation at Asn34. It is decelerated the more the size of the carbohydrate moiety increases. However, it was possible to estimate the thermal unfolding constants by extrapolation of the kproteolysis-values to infinite protease concentrations. Here, the steric shielding effects were negligible and it was found that all the non-RNase A species possess nearly the same unfolding rate constants and consequently the same kinetic thermal stability. This means that the stability difference between RNase A and RNase B has to be attributed to the attachment of the first carbohydrate unit to Asn34. According to the computer modelling studies by Woods et al. (1994), a hydrogen bond between the side chain of Lys37 and GlcNAcl might be responsible for the stabilization effect of the glycosylation in RNase B.
126 ACKNOWLEDGEMENTS
We thank Dr. A. Schierhom, Martin-Luther University, Halle, for performing MALDI mass spectrometry measurements and the Deutsche Forschungsgemeinschaft, Bonn, Germany, for supporting this work. REFERENCES
Arnold, U., Rticknagel, K. P., Schierhorn, A. & Ulbrich-Hofmann, R. (1996) Thermal unfolding and proteolytic susceptibility of ribonuclease A, Eur. J. Biochem. 237, 862869. Arnold, U. & Ulbrich-Hofmann, R. (1997) Kinetic and thermodynamic thermal stabilities of ribonuclease A and ribonuclease B, Biochemistry 36, 2166-2172. Arnold, U., Schierhorn, A. & Ulbrich-Hofmann, R. (1998) Influence of the carbohydrate moiety on the proteolytic cleavage sites in ribonuclease B, J. Prot. Chem. /7, 397-405. Berman, E., Wakers, D. E. & Alerhand, A. (1981) Structure and dynamic behavior of the oligosaccharide side chain of bovine pancreatic ribonuclease B, ]. Biol. Chem. 256, 3853-3857. Blum, H., Beier, H. & Gross, J. H. (1987) Improved silver staining of plant proteins, RNA and DNA in polyacrylamide gels, Electrophoresis 8, 93-99. Catanzano, F., Graziano, G., Capasso, S. & Barone, G. (1997) Thermodynamic analysis of the effect of selective monodeamidation at asparagine67 in ribonuclease A, Prot. Sci. 6, 1682-1693. Imoto, T., Yamada, H. & Ueda, T. (1986) Unfolding rates of globular proteins determined by kinetics of proteolysis, J. Mol. Biol. 190, 647-649. Jaenicke, R. (1991) Protein stability and molecular adaptation to extreme conditions, Eur. J. Biochem. 202, 715-718. Lis, H. & Sharon, N. (1993) Protein glycosylation. Structural and functional aspects, Eur. ]. Biochem. 218,1-27. Rademacher, T. W., Parekh, R. B. & Dwek, R. A. (1988) Glycobiology, Ann. Rev. Biochem. 57, 785-838. Rudd, P. M., Joao, H. C., Coghill, E., Fiten, P., Saunders, M. R., Opdenakker, G. & Dwek, R. A. (1994) Glycoforms modify the dynamic stability and functional activity of an enzyme, Biochemistry 33, 17-22. Sch~igger, H. & von Jagow, G. (1987) Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from I to 100 kDa, Anal. Biochem. 166, 368-379. Williams, R. L., Greene, S. M. & McPherson, A. (1987) The crystal structure of ribonuclease B at 2.5-A resolution, ]. Biol. Chem. 262,16020-16031.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
127
Comparative studies on selective modification of E-amino groups in lipases and phospholipase A2 Bart C. Koops, Arend J. Slotboom and I-Iubertus M. Verheij Department of Enzymology and Protein Engineering, C.B.L.E., Utrecht University, PO Box 80054, 3508 TB Utrecht, The Netherlands Several lipases and phospholipase A2 (PLA2) were chemically modified with ethyl caprylimidate, the capryl ester of p-hydroxyphenyldimethylsulfonium methyl sulphate (CsDSP-ester), the capryl carbonate of p-hydroxyphenyldimethylsulfonium methyl sulphate (C8DSP-carbonate) and polyethylene glycol monomethyl ether activated with p-nitrophenyl chloroformate (pNP-MPEG). PLA2 could be easily modified to high degrees, whereas modification of lipases was less efficient. The poor modification with the C8-DSP-ester, C8DSP-carbonate, and pNP-MPEG can be explained by hydrolysis of the reagents by the lipases and by impurities present in the crude lipase preparations. Purification of the lipases decreases the hydrolysis rate of the reagent, but the enzymatic hydrolysis of the reagent remains considerable. For purified Candida rugosa lipase the hydrolysis of pNP-MPEG is slow enough to give a relatively high degree of modification when reacted with this reagent. The degree of modification for Candida antarctica lipase B and Staphylococcus hyicus lipases could be enhanced by increasing the pH of the modification reaction and addition of the reversible inhibitor, 2,4-dichlorobenzeneboronic acid. 1. I N T R O D U C T I O N During the past decade, the use of enzymes as biocatalysts has been established as an important tool in organic synthesis. Due to their ability to discriminate between enantiomers they can be utilised as biocatalysts for the kinetic resolution of racemates to produce chiral building blocks for pharmaceuticals and fine chemicals. Furthermore, hydrolytic enzymes can catalyse the reverse reaction in organic solvents. By far the most frequently used biocatalysts in organic synthesis are lipases (triacylglycerol hydrolases, EC 3.1.1.3) because they are relatively cheap, available from many sources and have a broad substrate selectivity. However, the use of lipases in organic synthesis on industrial scale has two major disadvantages. First, enzymes are insoluble in non-aqueous solvents whereas most of the reactants are soluble in these media. For such heterogeneous reactions the activity of the lipase is low and therefore a large amount of the biocatalyst is needed. Second, their stability under conditions used in industrial applications is rather low and the biocatalyst can only be used for a short period of time. These disadvantages make the use of lipases as biocatalyst in industrial processes approximately ten times too expensive. Obviously improving the performance of enzymes in organic media should concern their activity, solubility and stability in these media. An often described strategy is changing the surface properties of the enzymes by covalent coupling of hydrophobic or amphiphilic groups to the e-amino group of lysine residues. Most papers on this subject describe either reductive alkylation with aldehydes (1), amidination with imidates (2), acylation with esters of phydroxyphenyldimethylsulfonium methyl sulphate (DSP-esters) (3) or the introduction of a polyethylene glycol moiety (pegilation) with activated polyethylene glycol monomethyl ethers (MPEG) of different molecular weights (4, 5). All these publications describe an increased activity and thermal or solvent stability in organic media. However, only for pegilated enzymes
128 an increase in the solubility in organic solvents has been described. MPEG was previously introduced by activation with (toxic) cyanuric chloride. As an alternative, milder reagents for pegilation are used, such as MPEG activated with p-nitrophenyl chloroformate (pNP-MPEG) (6), or succinimidyl succinate (suc-MPEG) (7) There are numerous articles on the topic of chemical modification of enzymes, but only a limited number of lipases have been subjected to modification. In this article we describe the modification of several lipases with the four modifying agents depicted in Figure 1. All these reagents react with the E-amino group of lysines and with the m-amino group. Amidination of proteins with imidoesters results in retention of the positive charge at the lysines. Modification of proteins with the DSP-ester, DSP-carbonate, and pNP-MPEG results in a neutral amide linkage for the first or carbamate linkage for the latter two reagents.
J H
100
+ Cl-
C7H15
~8o
O--C2Hs /
,CH3
Xc.~
CH3SO4-
C7Hls-O--CO--O-CsH4-(CH3 CHaSO4CH3
4
-
C
.o_
C7H1s --CO-O--CsH4-S./+
3
100
CH3-(OCH2CH2)110--O--CO--O-CsH4-NO2
Figure 1. Structures of the modification reagents. 1. Ethyl caprylimidate. 2. CSDSP-ester 3. Cs-DSP-carbonate 4. pNPMPEG5000. C6H4 is para substituted phenyl.
80
~
..'g =_60
60 =o ,..e ~.
"~ 40
40 o~
~
"O
e
20 0
20
0
,,,J,,,I,,,i,,,i 20 40 60 molar excess
80
,,,
0 100
Figure 2. Degree of amidination and activity of PLA2 as a function of the molar excess of ethyl caprylimidate. Degree of amidination determined with ninhydrin (o) or with amino acid analysis (e). Residual activity after modification (m)
2. MATERIALS AND METHODS 2.1. Enzymes Candida antarctica lipase B, SP 525 (CALB) and Rhizomucor miehei lipase, Novozym 388 (RML) were kind gifts of Novo Nordisk (Bagsvaerd, Denmark). Cutinase from Fusarium solani pisi and Pseudomonas glumae lipase (PGL) were generously donated by Prof. Dr. M.R. Egmond (Unilever R.L., Vlaardingen, The Netherlands). Candida rugosa lipase, Lipase MY (Meito Songyo, Japan) (CRL) and Pseudomonas fluorescence lipase, Amano P (PFL) were generous gifts of Dr. G.J.M. van Scharrenburg (Solvay-Duphar, Weesp, The Netherlands). Bacillus subtilis lipase (BSL) was a kind gift from Dr K. Schanck-Brodrtick from Universit6 Catholique De Louvain (Louvain-la-neuve Belgium). Geotrichum candidum lipase (GCL) was obtained from Biocatalysts Ltd (England). Porcine pancreatic lipase (PPL) was obtained from Calbiochem (Los Angeles, USA). Porcine pancreatic phospholipase (PLA2) was obtained as described previously (8). Staphylococcus hyicus lipase (SHL) was isolated as described before (9). 2.2. Chemicals Ethyl chloroformate was from Acros Chimica. 4-(Methylmercapto)phenol was obtained from ICN Biomedicals. Dimethyl sulphate, octyl chloroformate and p-nitrophenyl
129 chloroformate were purchased from Aldrich. 2.4-Dichlorobenzeneboronic acid was obtained from Johnson Matthey. Sodium deoxycholate (DOC) was from Merck. Po' ",~thylene glycol monomethyl ether (MW 5000) was purchased from Fluka. Sigma lipase sabstrate (stabilised olive oil emulsion) was obtained from Sigma. All other chemicals were of analytical grade. Ethyl caprylimidate, C8-DSP-ester, and pNP-MPEG5000 were synthesised in analogy to the procedures described by Suydam et al. (10), Kouge et al. (11) and Veronese et al. (6), respectively. The C8-DSP-carbonate was synthesised analogously to the Cs-DSP-ester. Thin-layer chromatography was done on TLC plates from Merck (silicagel DC Fertig). 1HNMR measurements were done on a 300 MHz Brucker or Varian machine. Mass spectrometry data was obtained on a JMS SX 102/102 FAB Spectrometer. Small-scale gel filtration PD-10 columns (Sephadex G-25) were purchased from Pharmacia.
2.3. Preparation and partial purification of the crude lipases All lipases except BSL and RML were dissolved in distilled water, centrifuged to remove insoluble matter, dialysed against distilled water for three days at 40C and lyophilised prior to use (crude lipase preparations). Under these conditions no loss of enzymatic activity occurred. BSL was dialysed against 10 mM glycine buffer pH 10 to prevent precipitation. Because RML contains cellulases it was passed over a PD-10 column for desalting. Partial purification of CALB, RML and CRL was done on a DEAE-cellulose column equilibrated with 25 mM Tris.HCl pH 7.0, 7.5 and 7.0, respectively. BSL was purified on a CM-cellulose column equilibrated with 10 mM sodium borate pH 9.0. All purifications were carried out at 4~ Under these conditions CALB did not bind to the column. RML and CRL were eluted with a linear salt gradient (0 - 0.5 M NaCI). BSL was stepwise eluted with 0.1 M NaCI, 0.5 M NaC1 and finally with 10 mM NaC1 in 10 mM sodium borate at pH 12. All fractions with lipase activity were combined, dialysed against distilled water or 10 mM glycine buffer pH 10 and lyophilised. PPL was purified according to the procedure described by Verger et al. (12). After purification the CALB, RML, CRL and BSL preparations contained approximately 30-50% lipase (w/w) and PPL contained more than 90% lipase (w/w) based on SDS-PAGE.
2.4. Modification of PLA2 and lipases PLA2 was amidinated by the procedure adapted from Wofsy and Singer (13). Phospholipase A2 solutions (0.2%) in 0.1 M borate buffer at pH 8.5 were reacted with ethyl caprylimidate at different molar ratios of the reagent over the protein amino groups (Figure 2) at room temperature. The pH of the reaction mixture was re-adjusted to 8.5 with 2 M NaOH. After 4 hours the remaining lipolytic activity was measured and the reaction mixture was passed twice over a PD-10 column to remove excess reagent and hydrolysis products. The degree of modification was determined with ninhydrin. Alternatively, the reaction mixtures were dialysed 16 hours against 50% DMF, 24 hours against distilled water and prepared for amino acid analysis. The modified enzyme preparation was stored at -20~ The lipases were modified similarly, except that after modification the excess reagent was not removed. RML was amidinated at 0~ to prevent proteolytic degradation. Phospholipase A2 and lipases were acylated with the C8-DSP-ester and Cs-DSP-carbonate according to the procedure of Kawasaki et al. (14) with minor changes. To 900 lxl of the protein solution (0.56 mg/ml in 50 mM pyrophosphate buffer, pH 8.0), 100 ~tl of the DSP-ester (or DSP-carbonate), dissolved in the same buffer, was added at 40C. Based on the concentration of the amino groups, different molar excesses of the reagent were added (Table 2). After 24 hours for the DSP-ester modification, and 3 hours for the DSP-carbonate modification the degree of acylation was determined with ninhydrin. The proteins were pegilated as described by Veronese et al. (6). Protein solutions (1 mg/ml) in 50 mM pyrophosphate buffer at pH 8.0 or 50 mM borate buffer pH 8.8 were reacted with activated pNP-MPEG5000 at different molar ratios of the reagent over the protein amino groups (Table 2). After 24 hours reaction at 4~ the degree of modification was measured with ninhydrin. In case that a reversible inhibitor was used during the modification, 20 mM of 2,4dichlorobenzeneboronic acid was added to the lipase solution prior to addition of the reagent.
130
2.5. Determination of the degree of modification. The concentration of amino groups before and after modification was determined in triplicate with ninhydrin (15-17). Therefore, 50 gl of protein solution (1-2 mg/ml) was mixed in a 1.5 ml eppendorf vial with 100 gl distilled water, 50 gl 4 M acetate buffer pH 5.1 and 200 I.tl ninhydrin reagent. The mixture was vortexed and heated at 100*C for 15 minutes, cooled at 100C for 10 minutes and diluted with 1 ml 50% ethanol. After vortexing, the eppendorfs were centrifuged at 14000 rpm and the absorbance at 570 nm was measured against a blank. The extinction coefficient of ninhydrin was determined with a glycine solution of known concentration. The degree of modification was calculated as (1 - [NH2]after / [NH2]before). 100. Alternatively, the amidinated proteins were dialysed in 50 % DMF overnight followed by dialysis against distilled water for 24 hours in stead of the PD-10 column. The dialysed samples were hydrolysed in constant boiling HCI and analysed by amino acid analysis. In this case the degree of modification (%) was calculated as (1 - [lys]after/[lys]before).100.
2.6. Enzymatic activity measurements pH-Stat measurements: (Phospho)lipase activity was assayed at 40"C and pH 8.0 with a pHStat titration set (Radiometer, Copenhagen, Denmark) consisting of a TTTld pH meter, a SBR2c recorder and an ABU12 autoburette containing a 20 mM NaOH solution. Four assay conditions were used. (i) Tributyrin / Triton: the reaction mixture consisted of 120 mM Triton X100, 60 mM tributyrin, 100 mM NaCI, 5 mM CaC12 and 5 mM Tris.HC1. (ii) Tributyrin / DOC: the reaction mixture consisted of 110 mM tributyrin, 2 mM DOC, 150 mM NaC1 and 5 mM Tris.HCl. (iii) Sigma lipase substrate (stabilised olive oil emulsion 50% v/v): the reaction mixtul:e consisted of Sigma lipase substrate, 6 times diluted in 50 mM NaC1, 10 mM CaC12 and 5 mM Tris.HC1. (iv) Egg-yolk lecithin: the reaction mixture consisted of 20 ml 6 mM DOC plus 10 ml of the following mixture: one egg-yolk in 130 ml 20 mM CaC12. One (phospho)lipase unit is defined as the amount of enzyme which liberates 1 grnol of fatty acid per minute. Spectrophotometric assay: Lipase activity was assayed at room temperature with 250 gM acyl p-nitrophenyl esters (pNP-ester) in 100 mM Triton X100, 50 mM NaC1, 5 mM CaC12 and 5 mM Tris.HC1 at pH 8. The pNP-esters were dissolved in acetonitrile prior to mixing with the Triton solution (1% acetonitrile in the assay). For cutinase and CALB p-nitrophenyl butyrate was used as substrate, for SHL, CRL and RML p-nitrophenyl caprylate was used. To 1 ml substrate solution, 10 gl of lipase solution was added. The release of p-nitrophenoxide ion following lipase hydrolysis of the substrate is recorded at 400 nm using a Shimadzu UV 1205 spectrophotometer connected with a Shimadzu U-125MU recorder. The apparent molar extinction coefficient of the p-nitrophenoxide ion in the presence of 100 mM Triton X 100 was found to be 10.000 M-1.era-1. One lipase unit was defined as the amount of lipase which liberates 1 ~nol ofp-nitrophenoxide ion per minute. The rate of hydrolysis of the DSP-ester, DSP-carbonate and pNP-MPEG by lipases was measured in a similar way. The reaction mixture contained the reagent (150-300 ~IVI) in 50 mM pyrophosphate buffer (pH 8.0). Upon addition of lipase the release of DSP or p-nitrophenol was followed by measuring the increase in absorbance at 269 or 400 nm, respectively. The release of DSP or p-nitrophenol due to chemical modification is negligable since the amount of protein in the activity test is very low.
2.7. Inhibition of lipases by 2,4-dichlorobenzeneboronic acid. Lipase activity was measured with p-nitrophenyl esters as described above. In the assay 10 gl of the inhibitor (I) in methanol was added before adding the enzyme. The inhibitor concentration was varied between 100 nM and 1 raM. The reciprocal hydrolysis rates were plotted against the inhibitor concentration and the data were fitted to equation (1). The intercept with the x-axes represents the negative value of Kiapp.
• =
1 +
1 )
KM
1
"is] Vmax + (Vmax "IS]"
1 ).[I]
(1)
131 3. R E S U L T S
3.1. Amidination of PLA2 and lipases with ethyl caprylimidate PLA2 was reacted with ethyl caprylimidate in a molar excess (mol / mol amino group) ranging from 2 to 100. By taking samples at different time intervals and measuring the concentration of amino groups, we determined that after 4 hours the reaction reached a maximum (data not shown). The results of amidination of PLA2 are depicted in Figure 2. Obviously, increasing the molar excess of imidate results in an increase in amidinated amino groups with a maximum at 80%. The difference in the degree of amidination found with the ninhydrin method and amino acid analysis is probably due to selective loss of modified PLA2 during the purification of the reaction mixture on the PD-10 columns in the first method. It was found that PLA2 with a high degree of modification sticks better to the PD-10 column than PLA2 with a low degree of modification. The rapid decrease of the PLA2 activity with increasing amidination is not surprising, since modification of the o~-amino group inactivates PLA2 (18, 19). For the modification of crude lipase preparations it was assumed that the average reactivity of amino groups in different proteins would be similar. Therefore, the lipases were reacted with a 2, 25 and 50 fold molar excess of ethyl caprylimidate , to obtain lipases of which approximately 25, 50 and 75% of their amino groups were amidinated. To test this assumption we subjected the lipases modified with a 50 fold molar excess to amino acid analysis. It was found that 25-32% of the amino groups were amidinated (Table 1). While PLA2 lost 92% of its activity upon modification, most lipases remained active. 3.2. Modification of PLA2 and lipases with Cs-DSP-ester,
and pNP-MPEGsooo.
Cs-DSP-carbonate
PLA2 was modified with Cs-DSP-ester, C8-DSP-carbonate and pNP-MPEGs000 at 4~ During 48 hours samples were taken to determine the degree of modification. For the modification reaction of the C8-DSP-ester and pNP-MPEGs000, no increase in the degree of modification was found after 24 hours. For the modification with Cs-DSP-carbonate the reaction reached a maximum after 3 hours. Obviously, an increase in the molar ratio of reagent over protein amino groups resulted in an increase in the degree of modification (Table 2). As was found for the amidination of PLA2, the activity decreased with increasing acylation in a similar way (data not shown). Table 1. Degree of amidination and lipase activity before and after modification with a 50 fold molar excess of ethyl caprylimidate. activity Enzyme amidination e native enzyme .... amidinated enzyme (%) (O/m~) (U/ml~) PLA2 79 1200d 96d CRL 29 39 b 6b GCL _f 27 c 1c CALB 25 181 b 200 b PPL 28 2470 b 552 b PFL _f 63 a 40 a PGL 32 458 a 314 a RML _f 98 c 82 e BSL 27 8b 5b cutinase 32 532 a 236 a a Tributyrin/Triton assay, b Tributyrin/DOC assay, c Sigma lipase substrate, d Egg-yolk assay. e The degree of amidination was measured with amino acid analysis, f The concentration of lysine after modification could not be determined accurately
132 Table 2. Degree of modification of PLA2 and (purified) lipases with Cs-DSP-ester, Cs-DSPcarbonate and pNP-MPEG. The number of lysine residues for each enzyme molecule are: PLA2: 9, CRL: +_21, PPL: 22, CALB: 9, RML: 7, SHL: 32 and cutinase: 6. Enzyme molar excess Degree of modification (%) Cs-DSP-ester Cs-DSP-carbonate pNP-MPEG50o0 PLA2 2 16 17 n.d. PLA2 5 30 47 44 PLA2 10 65 n.d. n.d. PLA2 20 79 n.d. 79 CRL* 5 0 0 n.d. CRL* 20 0 0 n.d. CRL 20 4 n.d. 48 PPL* 5 6 9 n.d. PPL* 20 8 11 n.d. PPL 20 35 n.d. n.d. CALB* 5 0 2 n.d. CAI.~* 20 10 12 n.d. RML 20 6 n.d. n.d. SHL 5 0 0 4 SHL 20 0 0 15 cutinase 5 n.d. n.d. 1 cutinase 20 n.d. n.d. 10 * Crude lipase preparations, n.d. = not determined. Subsequently, lipases were modified with the three reagents under the same conditions as PLA2. Compared to PLA2, the same molar ratio of reagent over amino groups resulted in lower degrees of modification (Table 2). A possible reason for this lower degree of lipase modification could be that the lipases are able to hydrolyse the modifying reagents or that impurities in the crude lipase preparation destroy the reagent. Therefore, we measured the hydrolytic activity of the lipases towards these reagents before and after partial purification. We found that after purification the rate of hydrolysis was lower, but that the partially purified lipases were still able to hydrolyse the reagents (Table 3). Modification of partially purified CRL and PPL with the Cs-DSP-ester resulted in a low (4%) and substantial (27%) increase in the degree of modification. The hydrolysis rates of some lipases presented in Table 3 seem very low but under the conditions used for modification these activities are too high. CRL for example, with an activity of 0.2 U/mg towards the Cs-DSP-ester, degrades 50% of a 20 fold molar excess of this reagent in 10 minutes. Pegilation of CRL, which has a low rate of hydrolysis towards pNP-MPEGs000, resulted in the highest degree of modification (48%). We conclude that the chemical modification of most lipases is severely hampered by the capability of the lipases to hydrolyse the reagent before modification can take place. Table 3. Hydrolysis rates of partially purified lipases towards the different reagents and Kiapp values of. 2,4-dichlorobenzeneboronic acid for the lipases. Lipase hydrolysis rate Kiapp .... Cs-DSP-'ester Cs-DSP-carbonate 'pNP-MPEG5000 (U/mg) (U/mg) (U/mg) lxM cutinase 203 10 SHL 112 31 CALB 0.5 0.09 CRL 0.2 0.04 RML 0.3 0.02 The substrate concentrations were 270 lttM for the DSP-ester, and 210 l.tM for pNP-MPEG.
1" 210 0.01 2.5 3 5.5 0.006 180 0.02 1430 160 ~tM for the DSP-carbonate
133 In order to get higher degrees of modification, it is necessary to decrease the activity of the lipases and/or increase the rate of the modification reaction. Increasing the rate of the modification reaction can be done by increasing the pH since the e-amino group of lysine residues are less protonated (and thus more reactive) at a higher pH. However, the pH-range is limited because these reagents are unstable due to alkaline hydrolysis. To decrease the lipase activity, we (reversibly) inhibited several lipases by 2,4-dichlorobenzeneboronic acid and determined their Kiapp values. As can be seen in Table 3 both SHL and CALB have high affinities for the inhibitor. Modification of SHL with pNP-MPEG5000 in the presence of 2,4dichlorobenzeneboronic acid increased the degree of modification only from 15 to 22%. When SHL was modified in the presence of the inhibitor at pH 8.8 the degree of modification substantially increased to 42%. Modification of CALB with the same reagent at pH 8.8 in the presence of the inhibitor resulted in 48% modification. 4. D I S C U S S I O N For the modification of the crude lipase preparations with imidoesters an easy determination of the degree of amidination with ninhydrin is not possible. The reason is that the excess of the imidoester interferes with the ninhydrin reaction, and thus has to be removed. However, upon removing the excess imidoester on a PD-10 column, not only the reagent but other impurities (amino groups) present in the crude lipase preparation are removed as well. Since the degree of amidination is calculated from the difference in the amount of amino groups before and after modification, it is clear that with this strategy large errors are obtained. Our assumption that the results obtained for amidination of PLA2 could be extrapolated to lipases proved to be not entirely true, since lower degrees of amidination were found for the lipases. Modification of lipases with an extremely high molar excess (300 fold) does not lead to a higher degree of modification, most probably because the insoluble hydrolysis products of the imidoester give rise to practical problems. Modification of lipases with the DSP-ester, DSP-carbonate, and pNP-MPEG also gives lower degrees of modification than with PLA2. The reason for this difference is clear, since the lipases and impurities in the lipase preparation are able to hydrolyse the reagents. Even though partial purification of the lipases results in a lower rate of hydrolysis of the reagent, most lipases hydrolyse a substantial part of the reagents before the modification reaction can take place. The relatively high degree of pegilation of CRL in the absence of inhibitor, can be reached because of a low hydrolysis rate. Only the use of a reversible inhibitor in combination with an increase in the pH improved the degree of modification for SHL and CALB. For cutinase, RML and CRL, the affinity for the reversible inhibitor was too low to be used in this way. In literature, modification of lipases with a DSP-ester and pNP-MPEG has only been described for Phycomyces nites lipase (3, 20) and CRL (21-24), respectively. A recently described modification reagent polyethylene glycol succinimidate, which is the most reactive pegilation reagent under neutral conditions (7), has only been described for CRL. We suggest that modification of other lipases with this reagent has its limitations, since this activated ester may be hydrolysed by many lipases. Based on our results, we conclude that setting up a general strategy for lipase modification should be done with (at least partially) purified lipases. Moreover, either a reagent should be used that cannot be hydrolysed by the lipases, or the reaction should be carried out in the presence of a potent competitive inhibitor.
134 ACKNOWLEDGEMENTS
This research was funded by the Dutch Ministry of Economics through IOP-Katalyse. We would like to thank Ruud Dijkman for the synthesis of p-hydroxyphenyldimethylsulfonium methyl sulphate and its esters, Dr. J.J. Kettenes- van den Bosch and C. Versluis for recording the NMR and Mass spectra, and H. Ravestein and P. Overkamp for the amino acid analyses. REFERENCES
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o
Mol. Catal. B: Enzym. 3, (1997) 171-176. .
.
.
5. 0
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8. .
10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.
Basri, M., Ampon, K., Yunus, W. M. Z., Razak, C. N. A., and Salleh, A. B. J. Am. Oil Chem. Soc. 69, (1992) 579-583. Murakami, M., Kawasaki, Y., Kawanari, M., and Okai, H. J. Am. Oil Chem. Soc. 70, (1993) 571-574. Inada, Y., and Matsushima, A. Biocatalysis 3, (1990) 317-328. Matsushima, A., Kodera, Y., Hiroto, M., Nishimura, H., and Inada, Y. J. Mol. Catal. B: Enzym. 2, (1996) 1-17. Veronese, F. M., Largajolli, R., Boccfa, E., Benassi, C. A., and Schiavon, O. Appl. Biochem. Biotechnol. 11, (1985) 141-152. Bremen, U., and Gais, H. J. Tetrahedron-Asymmetry 7, (1996) 3063-3066. Nieuwenhuizen, W., Kunze, H., and de Haas, G. H. Methods Enzymol. 32B, (1974) 147-154. Simons, J.-W. F. A., Boots, J.-W. P., Slotboom, A. J., and Verheij, H. M. J. Mol. Catal. B: Enzym. 3, (1997) 13-23. Suydam, F. H., Greth, W. E., and Langerman, N. R. J. Org. Chem. 34, (1969) 292296. Kouge, K., Koizumi, T., Okai, H., and Kato, T. Bull. Chem. Soc. Jpn. 60, (1987) 2409-2418. Verger, R., de Haas, G. H., Sarda, L., and Desnuelle, P. Biochim. Biophys. Acta 188, (1969) 272-282. Wofsy, L., and Singer, S. J. Biochemistry 2, (1963) 104-116. Kawasaki, Y., Murakami, M., Dosako, S., Azuse, I., Nakamura, T., and Okai, H. Biosci. Biotechnol. Biochem. 56, (1992)441-444. Moore, S., and Stein, W. H. J. Biol. Chem. 176, (1948) 367-388. Moore, S., and Stein, W. H. J. Biol. Chem. 211, (1954) 907-913. Hirs, C. H. W. Methods Enzymol. 11, (1967) 325-329. Abita, J.-P., and Lazdunski, M. Eur. J. Biochem. 30, (1972) 37-47. Slotboom, A. J., and de Haas, G. H. Biochemistry 14, (1975) 5394-5399. Takahashi, Y., Tanaka, K., Murakami, M., Kawanari, M., Kawasaki, Y., Tatsumi, K., and Okai, H. Biosci. Biotechnol. Biochem. 59, (1995) 809-812. Basil, M., Salleh, A. B., Ampon, K., Yunus, W. M. Z., and Razak, C. N. A. Biocatalysis 4, (1991) 313-317. Basil, M., Ampon, K., Wan Yunus, W. M. Z., Razak, C. N. A., and Salleh, A. B. J. Chem. Technol. Biotechnol. 64, (1995) 10-16. Hem~iz, M. J., S~nchez-Montero, J. M., and Sinisterra, J. V. Biotechnol. Tech. 10, (1997) 917-922. Hem~iiz, M. J., S~chez-Montero, J. M., and Sinisterra, J. V. Biotechnol. Bioeng. 55, (1997) 252-260.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.U Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
r
135
Analysis of the thermal deactivation kinetics of modified by chemoenzymatic glycosylation Maria Asunci6n LONGO ~ and Didier COMBES b
"Department of Chemical Engineering, University of Vigo, Lagoas-Marcosende, 36200 Vigo, Spain. bDrpartement de Grnie Biochimique et Alimentaire, UA-CNRS 544, Institut National des Sciences Appliqures, Complexe Scientifique de Rangueil, 31077 Toulouse Crdex, France.
I. INTRODUCTION Enzyme stability is a crucial factor to determine whether application of biocatalysis will be commercially successful. Catalytic proteins loose part of their activity when they are subjected to the action of heat, extreme pH or proteases. During the last decades, much research has focused on the improvement of enzymes behaviour in the conditions in which they were to be used, and especially on the increase of their thermal stability. The production of heat-resistant enzymes would allow carrying out enzymatic reactions at higher temperatures, and therefore, increasing conversion rates and substrates solubility and reducing the risk of microbial growth and the viscosity of the reaction medium [1]. Several strategies have been proposed: use of soluble additives [2], immobilisation [3], protein engineering [4] and chemical modification [1, 5]. The covalent binding of appropriate modifiers to the enzyme's surface residues appears as the most popular approach, and very interesting results have been reported. Protein surface hydrophilic/hydrophobic balance seems to play a major role on enzyme thermal stability [6]. Non polar aminoacids are organised on protein surface as hydrophobic clusters, whose distribution may determine properties such as solubility, activity, specificity and stability towards pH, temperature or proteolysis [ 1]. Thus, the modification of protein surface characteristics by chemical binding of modifiers appears as a good strategy to improve biocatalyst performance. Among the investigated modifiers, polysaccharides are outstanding for the improvements in enzyme properties achieved. [7, 8]. Several strategies may be used to assess enzyme thermal stability [9]. Among them, determination of half-life time at a certain temperature is often chosen, since it provides information about enzyme residual activity, an important parameter regarding industrial applications of biocatalysts. Nevertheless, when the effects of enzyme surface modification on thermal stability are investigated, a detailed study of enzyme deactivation kinetics could be necessary. Some enzymes show exponential decay of activity with time, while others present different non-exponential kinetics [10]. A number of models have been proposed to explain the mechanism of enzyme thermal deactivation [ 10-12].
136 In the present work, a-chymotrypsin has been modified by attachment of glycosidic chains of various lengths. This modification has been carried out by a procedure combining chemical and enzymatic reactions [13], and it induced a global increase in the surface hydrophilic character of the protein. The effects of enzyme surface glycosylation on thermal stability in aqueous environment have been investigated, considering both the variation in halflife times, and the study of the deactivation kinetics. So, both half-life times and kinetic and thermodynamic deactivation parameters could be compared, in order to verify whether the results were coincident or not. In some cases, the deactivation parameters pointed out certain amelioration in enzyme thermostability during one step of the deactivation mechanism, even if the observation of half-life times had indicated a decrease in thermostability.
2. EXPERIMENTAL
2.1. Material Bovine pancreas a-chymotrypsin (EC 3.4.21.1), CNBr and N-acetyl-L-tyrosine ethyl ester (ATEE) were obtained from Sigma. Radioactive labelled sucrose was from HEN Research Products. Levansucrase from Bacillus subtilis (EC 2.4.1.10) was kindly provided by Eridania Beghin-Say. All other reagents were analytical grade. 2.2. Assay of a-chymotrypsin hydrolytic activity The substrate utilised for assay of a-chymotrypsin hydrolytic activity was N-acetyl-Ltyrosine ethyl ester (ATEE), as previously reported [14]. 2.3. Chemoenzymatic glycosylation of a-chymotrypsin Glycosylation of the enzyme was carried out by a two step chemoenzymatie technique previously developed [ 13]. This method involves chemical binding of sucrose molecules to the enzyme surface in a first step, followed by a second step in which a glycosyltransferase transfers fructose residues from a donor to the enzyme-bound glycosidic moieties, lengthening them in a progressive way. The levansucrase from B. subtilis has been utilised as glycosidic chain lengthening enzyme. a-chymotrypsin in control solutions and samples was separated from residual reagents by anion exchange chromatography, using a Mono-Q HR 5/5 column connected to an FPLC (Fast Protein Liquid Chromatography) system, both from Pharmacia LKB. The column was eluted at 1 ml/min with a gradient of 0.02 M piperazine buffer pH 9.5 as eluent A and the same buffer containing 1 M NaCI as eluent B. Protein fractions were collected, dialysed and freeze-dried. Glycosylation degrees were determined by radioactive labelling [ 13]. 2.4. Determination of surface hydrophilic character of the enzymes Native and modified enzymes hydrophilicity was evaluated from the reversed phase chromatography profiles obtained in an HPLC Hewlett Packard 1090 equipped with a Nucleosil C18 5 ttm column (220x2.1 nun, SFCC). Following sample injection, the column was eluted at 0.5 ml/min with a gradient of H20/0.1% TFA as eluent A and ethanol/0.1% TFA as eluent B. After a 5 min step of 100% A, a linear gradient from 40% to 100% B in 15 rain
137 was applied. Eluted proteins were evaluated from the on-line measurement of the optical density at 280 nm. 2.5.
Thermal stability of the enzymes
2.5.1. Enzyme thermal deactivation curves Samples and controls were diluted to the desired concentration (5 g/l) in 10.4 M HCI, in order to minimise the risk of proteolysis. Tight-closed tubes containing 0.5-ml aliquots were placed in a water bath at 50~ taken out at regular time intervals and cooled down in ice. The residual activities were measured and referred to the initial values. 2.5.2. Determination of enzyme deactivation kinetic parameters The experimental enzyme deactivation data were fitted to modelled theoretical curves using a non-linear regression procedure based on the Marquardt-Levenberg method of iterative convergence included into the Sigmaplot 1.02 (1994) software. All the relevant deactivation parameters were calculated.
3. RESULTS AND DISCUSSION The well-known bioeatalyst a-chymotrypsin has been modified, using a chemoenzymatic glycosylation method, in order to produce several enzyme forms glycosylated at various degrees. The extent of sugar binding achieved [13] ranged from 12 to 85 moles of grafted monosaccharide per mol of enzyme (0.08 to 0.55 g g'~). The hydrophilic/hydrophobic balance of the produceA enzymes was assessed from their reversed phase chromatography profiles. Attachment of glycosides brought about a decrease in elution times, which could be related to a progressive increase in the global hydrophilic character of the biocatalyst with glyeosylation. Then, thermal stability of some selected modified enzymes was thoroughly investigated, and compared to that of the native form. The evaluation of half-life times (Table 1) showed a very slight increase in thermostability for the less glycosylated forms (half-life times 5-10% higher than the control), and a decrease for the others, a-chymotrypsin bound to 19 moles of monosaccharide per mol of enzyme (0.123 g.g'~) showed a half-life time 28% lower than the non-modified form. When higher glycosylation degrees were obtained, half-life times decreased remarkably (up to 80% in some cases). Nevertheless, a more detailed study of the thermal denaturation patterns for glycosylated a-chymotrypsin provided further information. The deactivation curves indicated a variable fractional rate of deactivation, and therefore, pointed out that more than one type of bond must be broken before a total loss of activity occurs (Figure 1). The results were fitted to a series-type enzyme deactivation model, which involves two first-order steps with one active precursor (El) and a final enzyme state (E2) with possible nonzero activity [ 15]. E
kl .~ E am
l~ .~Ea2
138
This model assumes that E, Et and E2 are enzymatic homogeneous states, with several specific activities, k~ and k2 are the first order deactivation rate constants (min"t) for each step, and oh and a2 are the ratio of specific activities EriE and E2/E respectively. Residual enzyme activity (a) could be expressed as a function of time by the following equation:
a~k~ k2-k;l a,k~ exP("k~t)-(lo~k:~)(czl--m)exp(-k2t)+
a = 1+ l ~ - k ,
~2
Parameters at, kt and k2 for native and modified a-chymotrypsin have been calculated from the experimental data, using a non linear regression method based on a MarquardtLevenberg iterative convergence algorithm. The ratio a2 was considered equal to 0, which means that the final form of the enzyme is totally deactivated [ lO, 15].
- 1,0 0,8
A
1,0
0,6
.~ 0,8
0,4 ~~
"~t
"~176
.-~ 0,6
9
"~176
0,4 i
I
I
0
5
10
I
I
I
15 20 25 Time (min)
'i
I
30
0
5
I
'I
'
I
I
10 15 20 25 Time (rain)
0,2
~7 I
0,1
30
Figure 1. Thermal denaturation of glycosylated a-chymotrypsin A. Increase in half-life times. B. Decrease in half-life times. Moles ofmonosaccharide per mol of enzyme: Q 0 ; A 12; V 19; ~7 45
The thermodynamic parameter standard free energy (AG,~ of the thermal deactivation process has also been calculated for each step of the mechanism, from the equation: -AG,~ RT In [keTCh]
139 where ks is Boltzmann's constant (J K'l), h is Planck's constant (J min), K is the gas constant (J mol ~ K'~),T is the temperature (K) and ki is the deactivation rate constant (min'~). The values of the kinetic and thermodynamic parameters of deactivation are shown in Table 1. Table 1 Half-life times, kinetic and thermodynamic parameters of deactivation for glycosylated achymotrypsin (5 g/l solution in 104 M HCI, 50~ .....
,
Monosaccharide per
Half-life time
enzyme (mol mol"~)
(min)
0
16.2
12
otl
kl
k2
AG~
(min"l)
(min'b
0d morb
0.318
0.0814
3.04x10 "f~
246.3
17.2
0.420
0.115
9.0x10 "l~
242.4
19
11.6
0.412
0.164
2.08x10 "~
251.6
45
1.5
0.277
0.838
0,171
185.9
In some cases, the evolution of thermal deactivation after glycosylation, as indicated by half-life times did not totally agree with the values of the kinetic and thermodynamic parameters of deactivation obtained. For instance, binding of or to 12 moles of monosaccharide per mol of enzyme brought about a slight increase in half-life time, but the deactivation rate constants k~ and k2 increased and the standard free energy decreased after glycosylation. The study of half-life times indicated a stabilisation of the protein after modification, while the thorough study of the deactivation kinetics pointed out, to a certain extent, the opposite effect. On the other hand, the enzyme bound to 19 moles of monosaccharide per tool of protein, whose half-life time decreased in 28% compared to the control, undergone a global stabilisation effect, according to the observed increase in AG~ (= 5 ld mol'~). Furthermore, an increase in the value of r (0.318 for the control and 0.412 for the glycosylatod enzyme) and a decrease in the deactivation rate constant k~, were also detected. The former would imply a higher specific activity of the intermediate form El, and the latter a stabilisation effect during the second step of the deactivation process, which would account for the contradictory trends indicated by half-life time and AG~ results. The other glycosylated form studied (45 moles of monosaccharide per tool of enzyme) showed similar tendencies in half-life times and kinetic and thermodynamic parameters of deactivation. When ot-chymotrypsin was bound to small amounts of glycosides, its thermostability remained nearly constant or underwent certain amelioration, while for higher glycosylation degrees, it decreased remarkably. The present work seems to support the occurrence of a close relationship between the surface hydrophilic character and the thermal stability of enzymes. The mechanisms of thermal stabilisation of enzymes in aqueous environment have been thoroughly studied, and it seems
140 clear that the interactions between protein surface hydrophobic clusters and surrounding water play a key role in the heat-induced denaturation of biocatalysts [16]. Some researchers have proposed the defivatization of enzyme surface with hydrophilic molecules as a way to protect the non-polar clusters from contact to water, and therefore to increase thermal stability [1718]. The modifiers should trap the surrounding water molecules and create a protecting shield to the hydrophobic clusters. Nevertheless, it has to be considered that these hydrophilic modifiers could also attract water, and facilitate its contact with enzyme surface instead of protecting it against damaging interactions. This hypothesis would support the results presented here, in which the significant increase in hydrophilic character of the enzyme surface by glycosylation brought about a decrease in half-life times. Nevertheless, phenomena provoked by enzyme surface glycosylation appear to be quite complex. It is very difficult to know the protein sites to which glycoside molecules have been bound, and therefore the interactions induced. A thorough study of the enzyme deactivation kinetics will provide a more detailed information about the consequences of the modifications, and could reveal secondary stabilisation or destabilisation effects. In the present work, a stabilisation of the enzyme during the second step of the thermal deactivation process has been detected for some degrees of modification, in spite of the lowering in half-life times. This effect could be due to the binding of some molecules to sites in which advantageous interactions would occur.
REFERENCES 1. Mozhaev, V.V., Berezin, I.V. and Martinek, K. CRC Crit. Rev. Biochem., 173 (1988) 147. 2. Schmid, R.D. Adv. Biochem. Eng., 12 (1979) 41. 3. Monsan, P. and Combes, D. Methods Enzymol., 137 (1988) 584. 4. Gupta, M.N. Biotechnol. Appl. Biochem., 14 (1991) 1. 5. Roig, M. G. and Kennedy, J. F. CRC Cfit. Rev. Biotechnol., 12 (1992) 391. 6. Dill, K. A. Biochemistry, 29 (1990) 7133. 7. Lenders, J.P. and Crichton, R.R. Biotechnol. Bioeng., 26 (1984) 1343. 8. Marshall, J.J. TIBS, 3 (1978) 79. 9. Rupley, J. A., Gratton, E. and Careri, G. TIBS, 8 (1983) 18. 10. Henley, J.P. and Sadana, A. Enzyme Microb. Technol., 14 (1985) 42. 11. Lumry, R. and Eyring, H. J. Phys. Chem., 58 (1954) 110. 12. Gianfreda, L., Marrucci, G., Gfizzuti, N. and Greco, G. Biotech. Bioeng., 27 (1985) 877. 13. Longo, M.A. and Combes. D. J. Mol. Catal. B: Enzymatic, 2 (1997) 281. 14. Lozano, P., Combes, D. and lborra, J.L.J. Biotechnol., 35 (1994) 9. 15. Sadana, A. and Henley, J.P. Biotech. Bioeng., 30 (1987) 717. 16. Mozhaev, V. V. and Martinek, K. Enzyme Microb. Technol, 6 (1984) 50. 17. Mozhaev, V. V., Siksnis, V. A., Melik-Nubarov, N. S., Galkantaite, N. Z., Denis, G. J., Butkus, E. P., Zaslavsky, B. Yu., Mestechkina, M. and Martinek., K. Eur. J. Biochem., 173 (1988) 147. 18. Vankovfi, H., Pospisilov~, M., Tich~ M. and Tukovfi, J. Biotechnol. Techniques, 8 (1994) 375
StabilityandStabilizationof Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
141
Increasing the operational stability of flavoproteins by covalent cofactor binding Willem J. H. van Berkel, Robert H. H. van den Heuvel, Marco W. Fraaije and Colja Laane* D e p a r t m e n t of Biomolecular Sciences, Laboratory of Biochemistry, Wageningen Agricultural University, Dreijenlaan 3, 6703 HA Wageningen, The Netherlands
The active site residue Asp170 of the covalent flavoprotein vanillylalcohol oxidase was selectively changed by site-directed mutagenesis. Glu170 and Ser170 replacements resulted in flavinylated vanillyl-alcohol oxidase variants. However, substitution of Asp170 by Asn prevented covalent flavinylation without affecting flavin binding. The creation of flavindissociable mutant enzymes provides challenging possibilities to address the role of covalent flavinylation in the operational stability of vanillyl-alcohol oxidase.
1.
INTRODUCTION
Vanillyl-alcohol oxidase (VAO, EC 1.1.3.13) from Penicillium simplicissimum is a covalent flavoprotein acting on a wide range of lignin-derived phenolic c o m p o u n d s (1). VAO-mediated reactions of biotechnological relevance include the production of vanillin from creosol, the conversion of eugenol to coniferyl alcohol (2), and the enantioselective hydroxylation of short-chain 4-alkylphenols (3). The crystal structure of VAO, the first structure of a histidyl-FAD containing flavoenzyme (4), and sequence alignments suggest that VAO belongs to a new family of structurally related flavoenzymes (5,6). Many members of this family contain a histidyl-flavin, a feature shared by less than 5 % of all flavoproteins.
* This research was performed within the framework of the Innovation Oriented Research Program (IOP) Catalysis of the Dutch Ministry of Economy Affairs (project no. IKA96005).
142 The rationale for covalent flavinylation is unclear as is the mechanism by which the covalent bond is formed (7). As the efficiency of commercial flavoenzymes is often hampered by the dissociation of the flavin cofactor under stressful operational conditions, tailor-made covalent flavoenzymes could be of great value for biotechnological applications. Covalent bond formation may i) prevent inactivation of the cofactor, ii) improve resistance against proteolysis, and iii) improve protein stability. We have started to address the mechanism of covalent flavinylation by the characterization of VAO variants. The knowledge gained from these studies will be valuable for the future design of biotechnologically relevant flavoenzymes, made highly stable by the covalent linkage of the cofactor. Here we report on the preliminary characterization of Asp170 variants.
2.
2.1
MATE~ALS AND METHODS
Site-directed mutagenesis The vaoA gene has been cloned into pEMBL19(-) to give pIM3972 (5).
The plasmid pBC11, which is identical to pIM3972 except for a silent mutation at position 882, was used as vector for site-directed mutagenesis. The PstI-SalI fragment from pBC11 was cloned into pUCBM20 yielding a suitable construct for PCR-based mutagenesis. The oligonucleotide 5'-CTTGATGTACCGXXXCTTGGTGGCGGT-3' (where XXX denotes the replacement of GAT for GAG (D170E), TCT (D170S) and AAC (D170N), respectively) was used as primer for the construction of Asp170 variants. Mutated PstI-SalI fragments were excised from pUCBM20 and ligated into P s t I / S a l I digested pBC11. Succesful insertions were confirmed by plasmid sequencing.
2.2
Enzyme purification Enzyme purification was performed essentially as described previously (5). Highly pure Asp170 variants were obtained by introducing Superdex 200 PG 50/1000 gel filtration as final chromatography step. 2.3
Activity measurements Vanillyl-alcohol oxidase (VAO) activity was determined at 25 ~ in airsaturated 50 mM potassium phosphate, 1 mM vanillyl-alcohol, pH 7.5, by measuring the production of vanillin at 340 nm (1). 2.4
Gel electrophoresis SDS-PAGE was carried out as described (8). After electrophoresis the gel was incubated for 5 min in 4% acetic acid. Upon illumination with UV light, the fluorescence of protein-bound 8tx-(N3-histidyl)-FAD can be observed.
143
2.5
Analytical gel filtration
Analytical gel filtration was performed on a Superdex 200 HR 10/30 column using a Pharmacia .~kta system (9). 2.6
Guanidinium hydrochloride (GdmHC1) induced unfolding
Samples of VAO variants (2 ~M) were incubated at 25 ~ in 50 mM potassium phosphate buffer pH 7.5, containing 4 M GdmHC1. Fluorescence spectral analysis was done as described in (10).
3.
RESULTS AND DISCUSSION
3.1
General properties
The Asp170 variants were purified from E. coli TG2 in a three-column procedure, involving hydrophobic chromatography, hydroxyapatite chromatography and gel filtration. Analytical gel filtration established that all Asp170 variants were octameric and their yield after purification was comparable to that of the native recombinant enzyme (5). Illumination of unstained SDS-PAGE gels revealed that D170E and D170S were highly fluorescent, whereas no flavin fluorescence was detected with D170N. In line with this, treatment with 5% trichloroacetic acid yielded yellow protein precipitates for D170E and D170S and a yellow supernatant for D170N. This clearly demonstrates that the Asn170 replacement results in non-covalent flavin binding.
3.2
Catalytic properties
The Asp170 variants were less active with vanillyl-alcohol than the native enzyme. As can be seen from Table 1, almost no activity was observed with D170N. The highest activity was observed with D170E, t~ut the catalytic efficiency of this mutant was considerably decreased in comparison with the native enzyme (Table 1). These results suggest that Asp170 plays a crucial role in substrate conversion, which is in line with the proposed reaction mechanism (4).
144 Table 1 Kinetic parameters of VAO variants.
Enzyme
kc a t
Km
kcat / Km
s-1
~M
M-ls -1
VAO
3.3
160
20630
D170E
1.3
340
3820
D170S
0.01
nd
nd
D170N
< 0.005
nd
nd
nd, not determined
3.3
Flavin binding
D170N was eluted over a Biogel P-6DG column in 50 mM potassium phosphate, 2M urea, 1M KBr, pH 7.5. Under these conditions, and distinct from many other flavoproteins (11), no loss of flavin was observed. This shows that the covalent attachment of the flavin in vanillyl-alcohol oxidase is not essential for tight binding of the cofactor. 3.4
GdmHCl-unfolding
VAO activity is lost in 2 M GdmHC1 (12). When VAO was incubated in the presence of 4 M GdmHC1, a rapid increase in tryptophan fluorescence was observed (Fig. 1A). Moreover, the fluorescence emission maximum shifted from 340 to 355 nm, indicative of protein unfolding. Upon unfolding, no change in fluorescence intensity of p r o t e i n - b o u n d 80~-(N3-histidyl)-FAD was observed (inset Fig. 1A). This is consistent with the low fluorescence quantum yield of 8(x-(N3-histidyl)-FAD at neutral pH (13). When D170N was incubated in the presence of 4 M GdmHC1, a rapid shift in the tryptophan fluorescence emission maximum was followed by a slow increase in tryptophan fluoresence intensity (Fig. 1B). Unfolding of D170N was accompanied by a strong increase of flavin fluorescence (inset Fig. 1B). In analogy to the tryptophan fluorescence, flavin fluorescence increased slowly with time (inset Fig. 1B), suggesting that these secondary changes reflect flavin dissociation.
145
0.5
u.I
o
Z U.l 0 or) uJ IZ:
uJ o 0.4 z u.l
o.8
o
0.6 0.2
0
.-J u.
0.4
ill n"
0.2
-\ ,,
..i
~
n-
~,,,
~ L% %
320
~
450
,. -, _ _~
I
0
0
400
550
600
650
WAVELENGTH tnml
_
i
F ......
360
J
500
] 440
i
I
480
520
560
W A V E L E N G T H (nm)
1
o rr
S
0.5
0.8
~'~
0.6
,"
..',< ", \
/ ~
z 0.4
ix< \ /
,,,0.3
',,,',k
/,-., \
~
\
t
\
0.4 50
0.2 o 320
",
:i ~....._
50
WAVELENGTH (nm)
1
360
400
440
480
520
560
W A V E L E N G T H (nm)
Figure 1. Fluorescence emission spectra of VAO variants. Excitation w a s at 295 n m ( t r y p t o p h a n fluorescence) or 450 n m (flavin fluorescence, inset). (A) native VAO (---) and after incubation for 30 s in 4 M GdmHC1 (--). (B) D170 N (---) and after incubation for 30 s (- - -) or 30 min ( - - ) in 4 M GdmHC1.
146 4.
CONCLUSION
This is the first report on the properties of VAO variants. The results show that the generation of stable flavin-dissociable mutant enzymes is feasible and furthermore suggest that Asp170 is involved in activating the flavin ring for covalent attachment. Future protein-engineering will be directed towards the construction of highly active flavin-dissociable VAO variants. This will allow to study the operational stability of such enzymes in relation to their covalently-bound counterparts. REFERENCES
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.
12.
13.
M.W. Fraaije, C. Veeger and W.J.H. van Berkel, Eur. J. Biochem., 234 (1995) 271. W.J.H. van Berkel, M.W. Fraaije and E. de Jong, Process For Producing 4-Hydroxycinnamyl Alcohols, European Patent Application 0710289B1 (1997). F.P. Drijfhout, M.W. Fraaje, H. Jongejan, W.J.H. van Berkel and M.C.R. Franssen, Biotechnol. Bioeng., (1998) in press. A. Mattevi, M.W. Fraaije, A. Mozarelli, L. Olivi, A. Coda and W.J.H. van Berkel, Structure, 5 (1997) 907. J.A.E. Benen, P. S~_nchez-Torres, M.J.M. Wagemaker, M.W. Fraaije, W.J.H. van Berkel and J. Visser, J. Biol. Chem., 273 (1998) 7865. M.W. Fraaije, W.J.H. van Berkel, J.A.E. Benen, J. Visser and A. Mattevi, Trends Biochem. Sci., (1998) in press. M. Mewies, W.S. McIntire and N.S. Scrutton, Protein Sci., 7 (1998) 7. M.W. Fraaije, M. Pikkemaat and W.J.H. van Berkel, Appl. Environ. Microbiol., 63 (1997) 435. M.W. Fraaije, A. Mattevi and W.J.H. van Berkel, FEBS Lett., 402 (1997) 33. W.J.H. van Berkel, A.G. Regelink, J.J. Beintema and A. de Kok, Eur. J. Biochem., 202 (1991) 1049. F. Miiller and W.J.H. van Berkel, Chemistry and Biochemistry of Flavoenzymes Vol. I (F. Mfiller, ed.) CRC Press, Boca Raton, Florida (1991) 261. W.J.H. van Berkel, M.W. Fraaije, E. de Jong and J.A.M. de Bont, Flavins and Flavoproteins XI (K. Yagi, ed.) W. de Gruyter, Berlin, New York (1993) 799. E. de Jong, W.J.H. van Berkel, R.P. van der Zwan and J.A.M. de Bont, Eur. J. Biochem., 208 (1992) 651.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
147
Properties o f P E G - M o d i f i e d Microbial Proteases. Activity and Stability Studies
T. M. Fattma=, A. Agerlin Olsen', C. C. Fuglsang' and D. Otzenb
"Protein Discovery, Novo Nordisk A/S, Novo All6, 2880 Bagsvaerd, Denmark bDepartment of Biochemistry, University of Lurid, P.O Box 124, S-22100 Lund, Sweden
Attachment of PEG to enzymes has been reported to prevent (auto)proteolysis,' increase solubility and activity in organic solvents and to shield the enzyme from recognition by the immune system. 2 We now report results of activity and stability studies of PEG modified subtilisins. 1. Derivatization Two microbial subtilisins (Sub 1 and Sub2) were modified by attachment of succinimidyl carbonate activated mPEG 5000 in 80-100 times molar excess. The activated mPEG was coupled to the e-amino groups of the lysine side-chains to form stable urethane bonds. 3 0 CH3"{O
0 O--
0 +
NH2-Enzyme
~
CH3..,,O
O
O
n
NH-Enzyme +
HOO
2. Activity The catalytic activity of the PEG conjugates Table 1 towards peptide and protein substrates was Activity against Suc-AAPF-pNA tested and compared to the parent enzymes (figure 1). The Micha~lis Menten kinetic con(s") (M) (M"s "l) stants of the enzymes and derivatives against Subl 7.18E+05 1.160 6.19E+05 Suc-Ala-Ala-Pro-Phe-pNA are summarised in Subl-PEG 5.90E+05 1.052 5.61E+05 Table 1. The PEG-enzymes all have well preSub2 5.97E+05 0.369 1.62E+06 served activity, with l~t and K~ only slightly below the parent enzymes. The specific activ- Sub2?PEG 4.23E+05 0.338 1.25E+06 ity against a protein substrate (dimethyl casein) was typically 10-20% lower than the parent enzyme. This is probably due to steric hindrance near the active site. .
.
.
.
.
.
148
9 Sub1
* Subl-PEG 9 Sub2 9 Sub2-PEG
Activity
0
i--
0.0
I
I
I
i
I
0.5
1.0
1.5
2.0
2.5
IS]
Figure 1 Activity against Suc-AAPF-pNA 3. C o n f o r m a t i o n
To determine whether PEG induces any structural changes in the proteases, circular dichroism spectra of Sub 1 and Subl-PEG were obtained (figure 2). Both near UV and far UV spectra of the two proteins are identical, indicating that neither the tertiary nor the secondary structure are changed to any discernible extent by PEG conjugation. Far-UV CD-spectra
Near-UV CD-spectra 0.25
I
I
I
I
i
I
I
/
E 0.00
'7,
O "7,
O
E -0.25.
~
I
,=
u 25
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o
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--
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to
---Subl-PEG
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I
E
v
tO
-0.50
A 50
I
<3
-25 I
i
I
i
i
i
/
250 260 270 280 290 300 310 320 Wavelength (rim)
190 2()0 2';0 22.0 230 24,0 250 Wavelength (nm)
Figure 2. Near- and far-UV spectra of Subl and S u b l - m P E G in borate and succinic acid pH 6 with 1 mM CaCI 2 , pH 6.0. Buffer contribution subtracted. 4. Stability
The stability of the enzymes was studied by measuring the residual activity after incubation at 40~ for certain periods of time. Two different formulations were tested: buffer (5 mM succinic acid, 50 mM borate, 1 mM CaC12, pH 6.0) and buffer with 50% monopropyleneglycol (MPG). Aliquots were taken out and analysed for activity and for autolysis by HPLC. Results are illustrated at figure 3 where the legend 'Sub2 buffer' indicate parent Subtilisin2 dissolved in buffer, 'Sub2-PEG 50% MPG' indicate PEG-conjugated Subtilisin2 dissolved in buffer with 50% MPG, etc. In contrast to previously reported results, the PEG-proteases appeared to be less stable than the parent enzymes) The loss of proteolytic activity was followed by the
149
ActivitylA280 B0h Blh
1007
BB2h BB4h
75
E;~6h ITITn24 h
25 Sub2 buffer
Sub2 50% MPG
Sub2-PEGbuffer Sub2-PEG 50% MPG
Figure 3. Stability at 40~ The thermalstability of the enzymes was evaluated by differential scanning calorimetry (DSC). DSC analysis of Sub 1 and its PEG derivative is illustrated below (figure 4 and 5). Td = 73.3"C (PMSF-inhibited~
Co (call~ -0.001
Cp (cal/*C) .,
,o
-0.002
.~
No P"TU-q~
-0.003
-6 -1' ~ r PMSF-inhibited
-0.004 20
30
40
50
60
70
80
90
Temperature (*C)
CP (call*C) -0.004
,.0,
, 20
~ . 30
, 40
Sub2-mPEG5000 . . . . . , 9i 50
60
70
80
Temperature ('C)
Figure 6. DSC of Sub2 and Sub2-PEG
40
50 60 70 TemDerature ('C)
80
90
Figure 5. DSC of Subl-PEG +/- PMSF
Figure 4. DSC of Sub] +/- PMSF.
-0.010
30
9 90
While the parent enzyme showed only one peak at 70.9~ Sub 1-PEG revealed two peaks, one at 57~ and one at 71~ PMSF (Phenyl Methyl Sulfonyl Fluoride) inhibition does not alter the existence of two peaks, i.e. the first peak is not due to autolysis but rather to unfolding. Coupling of PEG to Sub2 also leads to two peaks but much closer to each other. The first peak appears at 71.4~ significantly higher than for Sub 1-PEG, thus conjugation does not seem to be so unfavourable for Sub2 as for Sub l (figure 6). This is reflected in the residual activity after incubation at 40~ (data not shown) where Sub2-PEG appear to be more stable than Sub 1- PEG.
150
5. Conclusion In contrast to the previously reported results, we found that the proteases were destabilised by attachment of PEG. The destabilisation showed as an increased susceptibility to autolysis at elevated temperatures. 6. Experimental Materials. The buffer used was 5 mM succinic acid, 50 mM borate, 1 mM CaC12, pH 6.0. Enzymes were subtilisins from alcalophilic Bacillus spp. All chemicals were purchased form Aldrich or Sigma. Preparation ofsuccinimidyl carbonate-PEG. Succinimidyl carbonate-PEG was prepared by reacting m-PEG with phosgene to obtain the chloroformiate, and treating this with N-hydroxysuccinimide following the procedure by Zalipsky et al. 3 Preparation of Subtilisin-PEG. Activated m-PEG 5000 (1.0 g ~ 0.20 mmoles) was dissolved in 0.1 N HCI (2ml) and added to the enzyme solution (50 mg enzyme ~ 0.0019 mmoles in 4 ml buffer). During the process, pH was maintained at 9-9.5 with 1 N NaOH. After stirring for 2 h at RT, pH was adjusted to 7 with 1 mM HC1. The reaction mixture was purified by gelfiltering on a Superdex 200 column eluted with 5 mM succinic acid, 50 mM borate, 200 mM NaCI, pH 6. Activity against Suc.-Ala-Ala-Pro-Phe-pNA. The catalytic activity was measured using a Cobas Fara analyzer. Assay buffer: 50mM borate, 150 mM KC1, lmM CaC12, 0.02% Brij35, pH 9.0. Enzyme concentration: 4.7 x 10"gM, substrate concentration: 0.03-2.1 raM, incubation temperature: 40~ Initial rates were calculated by following the release ofpnitroanilide at 405 nm (e = 7280 M "*cm "~) by direct fit to the Micha~lis Menten equation. Acknowledgement We wish to thank Jan Danielsen, Birgitte Ring Tote and Rie Kristine Schjeltved for technical assistance. REFERENCES 1. Yang, Z., Domach, M., Auger, R., Yang, F.X., Russel, A.J. (1996) Enz. Microb. Tehcnol. 18, 82-89 2. Inada, Y., Fur~awa, M., Sasaki, H., Kodera, Y. Hiroto, M., Nishimura, H., Matsushima, A. (1995) TIBTECH, 13, 86-91 3. Zalipski, S., Seltzer, R., Menon-Rudolph, S. (1992) Biotechnol. Appl. Biochem. 15, 100114
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
151
The effect o f crosslinking on thermal inactivation o f cellulases J. Bilen and U. Bakir Food Engineering Department, Middle East Technical University, 06531-Ankara, TURKEY
Cellulase complex was crosslinked by using three different crosslinkers, dimethyl suberimidate (DMS), dimethyl adipimidate (DMA) and dimethyl-3-3'-dithiobispropionimidate (DTBP) and their effects on thermal inactivation were investigated. These reagents are all homobifunctional bisimidoesters having different crosslinking distances and crosslink only amino groups to each other. No significant activity loss was observed due to crosslinking with none of the crosslinkers used. In general, DMA and DTBP crosslinking decreased the thermal stability slightly, however with DMS crosslinking, slight increases in thermal stabilities were observed.
1. INTRODUCTION Cellulose is the most abundant biological polymer on earth. It is primarily found as the major structural component of the cell walls of terretrial plants and marine algae but is also produced by other organisms such as some bacteria and marine animals. The rate of synthesis is about 7.5 x 101~ tons per year (1). Cellulose is the most promising renewable carbon source for a long range solution to resource problems of energy, chemicals and food. Cellulose can be hydrolyzed to soluble sugars either chemically or enzymatically. Enzymatic hydrolysis is becoming increasingly desired owing to mild processing conditions, high specificity of the reactions and better control of the process in many industrial applications. Cellulase is the enzyme complex used to degrade cellulose since it hydrolyzes cellulose to oligosaecharides and finally glucose. It contains at least three different components, endoglucanase (EC 3.2.1.4), cellobiohydrolase (EC 3.2.1.91) and 13-glucosidase (EC 3.2.1.21), and is produced by many microorganisms including Aspergillus, Fusarium and Trichoderma (2, 3).
* This research project (Misag-70) was supported by the Turkish Scientific and Technological Research Institute (TUBITAK).
152 The economics of an enzyme catalyzed reaction is mainly dependent on the lifetime of the catalitically efficient form of the enzyme under the reaction conditions employed. Therefore, stable enzymes or enzyme complexes, particularly thermostable enzymes, are the major concern in industrial applications. Many different techniques including protein engineering, immobilization and chemical modifications can be used to increase thermostability of enzymes (4, S). Chemical erosslinking is a chemical modification technique in which the compact structure is strengthen by bracing the protein intramolecularly or conjugating them intermolecularly to each other. Homobifunctional reagents carrying two identical functional groups have been demonstrated to be highly efficient in producing crosslinking between subunits of the macromolecules (6). Bisimidoesters react in alkaline solutions with amines to form amidines. Of the many reactive groups in proteins, only amino groups react and the products, like the amino groups they replace, are protonated at physiological pH (7).
2. MATERIALS AND METHODS
2.1. Materials Cellulase complex was kindly provided by NOVO Istanbul branch in Turkey. Avicel (micro crystalline cellulose) was bought from Merck AG. Carboxymethyl cellulose (CMC), bisimidoesters (DMS, DMA and DTBP) were obtained from Sigma Chem. Ltd., USA. All other chemicals used were analytical grade either from Merck or Sigma. 2.2. Cellulase Activity Measurements Cellulase was dissolved in 0.1 M acetate buffer, pH 4.8 and centrifuged at 10,000 x g at 4~ for 5 minutes in order to remove insolubles. Total cellulase activity was determined by using filter paper assay. 50 mg filter paper (Whatman No 1, I cm x 6cm), ruled as a small cylinder, was put into a test tube containing 1.5 ml enzyme solution, which was previously heated to 50~ The reactions were allowed to progress for 20 minutes and stopped by holding them in a boiling water bath for 2 minutes. Reducing sugar content as the result of hydrolysis was measured by Nelson-Somogyi method using glucose as a standard (8). Cellobiohydrolase activity was measured by using Avicel as a substrate. 1% Avicel solution was prepared in 0.1 M acetate buffer, pH 4.8. After addition of suitably diluted enzyme solution, it was incubated at 50~ for 20 minutes. To stop the reaction, the test tubes were kept in a boiling water bath for 2 minutes. After removing the residual Avicel by centrifugation for 3 minutes using a microcentfifuge at 12,500 rpm, reducing sugar content in the supernatant was analyzed by Nelson Somogyi method (8). Endoglucanase activity was measured by using CMC as a substrate. Substrate solution was prepared by dissolving 1% CMC in 0.1 M acetate buffer, pH 4.8. After mixing the substrate and enzyme solutions, the reaction mixture was incubated at 50 ~ for 15 minutes and stopped by addition of Nelson Somogyi reagent. Then, reducing sugar concentration was measured by Nelson-Somogyi method by using glucose as a standard (8). ff.glucosidase activity was measured by using eellobiose as the substrate at pH 4.8 and 37 ~ and the glucose formed due to the 13glucosidase activity was measured by using a glucose analyzer. One unit of enzyme activity was defined as the amount of enzyme required to release 1 ~tmol glucose equivalents per minute under the assay conditions for cellulase complex and its components. In all of the enzyme assays the reaction conditions were adjusted to measure initial activities.
153 2.3. Cross-linking Cellulase was dissolved in 0.1 M glycine-NaOH buffer at pH 9, centrifuged at 10,000 x g and 4~ for 5 minutes to remove insolubles. Cross-linkers (DMS, DMA or DTBP) were dissolved in the same buffer. After mixing the enzyme and cross-linker solutions at different concentrations, the reaction was allowed for 1 hour at room temperature. No termination step was considered due to the rapid self-hydrolyzation ofbisimidoesters (9). 2.4. Protein measurement Protein concentration was measured by a modified Lowry method (10). 2.5. Thermal inactivation experiments The pH of both the native and crosslinked enzymes were decreased to pH 4.8 in order to eliminate the pH effect during exposure to high temperatures. The temperatures of the enzyme solutions were increased to desired values quickly and after keeping them for certain periods, they were placed into an ice bath to decrease the temperature as soon as possible. Then, total cellulase, cellobiohydrolase and endoglucanase activities were determined as explained. All the experiments were at least duplicated and the average values were reported.
3. RESULTS AND DISCUSSION Since thermal inactivation usually starts with destruction of secondary and tertiary structure of the protein, the active conformation may be reinforced by chemical crosslinking. In this study, to stabilize the cellulase complex, three different crosslinkers were used and their effects on thermal inactivation were investigated. The crosslinkers used were dimethyl suberimidate (DMS), dimethyl adipimidate (DMA) and dimethyl-3-3'-dithiobispropionimidate (DTBP). These reagents are all homobifunctional bisimidoesters having different crosslinking distances, 9,11 and 13 A, respectively. Bisimidoesters were selected to crosslink cellulases since they crosslink only amino groups to each other, and no basic amino acid was reported to have an important function at the active sites of cellulases (7). Before starting the crosslinking experiments, pH stability of native cellulase complex was investigated to observe the stability of the complex at basic pH values required for the crosslinking reactions. No significant activity losses were observed even in 0.1 M pH 9 glycine buffer for three hours. Thermostability of the native cellulase complex was studied by incubating the enzyme complex at different temperatures from 55 to 75 ~ for 15 minutes and then calculating the residual activity of the complex and its components. As observed from Figure 1, cellobiohydrolase has the highest and 13-glucosidase has the lowest heat stability. In general, thermal inactivation starts in the range of 55 to 65 ~ In the crosslinking experiments, different crosslinker concentrations were used to find out the optimum crosslinker concentration to reduce the thermal inactivation of the complex. To observe the effect of crosslinking reaction on the activity of the enzyme system, the residual activities of total cellulase, endoglucanase and cellobiohydrolase were measured immediately after the crosslinking reactions. Measurements of 13-glucosidase activity was not performed due to the very low percentage (about 1 % ) of 13-glucosidase in the cellulase mixture. The results for the total cellulase activity are shown in Figure 2 for three crosslinkers used.
154 100
80 > o~,
u
,<
60
e=
"o .,..
rv.
40
20
25
55
60
65
70
75
Temperature,'C Figure 1. Heat Stability curves of native cellulase complex and its components. % Residual activities were calculated after 15 minutes of heat treatment. The data points were the averages of two separate experiments. (#: total cellulase activity, I1: cellobiohydrolase activity A: endoglucanase activity, x: 13-glucosidase activity)
Figure 2: The effect of crosslinking on the total cellulase activity. The data points were the averages of two separate experiments.
155 DMA crosslinking did not reduce either total cellulase or its component activities. With DMS and DTBP, the activity losses of cellobiohydrolases were slightly more than endoglucanases. However, total cellulase, endoglucanase or cellobiohydrolase activity losses due to crosslinking were not more than 10 %. No significant activity reduction after crosslinking probably indicates that no unfavorable interactions in the active sites were formed. In order to determine the optimum crosslinking concentrations for the detailed thermal inactivation studies, 15 minutes heat treatment at 67.5 ~ was applied to the native and crosslinked cellulases. Since DMA and DTBP crosslinked cellulases showed reduced thermostabilities than native cellulases at all the crosslinker concentrations used, no further thermal inactivation study was performed with these crosslinkers. However DMS crosslinked cellulases have about 10 % more residual activities than native cellulases. Since 0.2 % DMS concentration gave the maximum protection against heat, this concentration was determined as the optimum. The results of the thermal inactivation studies for total cellulase activity from 65 to 75 ~ are given in Figure 3.
(a)
._->" 10o .>
-<
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.
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.
.
.
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,
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(min)
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.
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Figure 3: The effect of DMS crosslinking on thermal inactivation of cellulases. % residual total cellulase activities are given at 65 ~ (a), 67.5 ~ (b), 70 ~ (c), 72.5 ~ (d) and 75 ~ (e). 0.2 % DMS was used. The data points were the averages of two separate experiments. ( | : crosslinked, e: native.)
156 DMS crosslinking seemed to increase thermostability at 65 and 70 ~ upon longer incubation periods, at 75 ~ crosslinking was not beneficial and better thermal stabilities were observed at 67.5 and 72.5 ~ Similar results were obtained for endoglucanase and cellobiohydrolase. Although DMS crosslinking seemed to change thermal inactivation, the changes were not very significant. REFERENCES
1. C.P. Kubicek, R. Messner, F. Gnaser, L.R. Mach, and M.E. Kubicek-Pranz, Enzyme Microb. Technol., 15 (1993) 90. 2. C.P. Kubicek, Adv. Biochem. Eng., 45 (1992) 1. 3. A. Singh and K. Hayashi, Advanc. Appl. Microb., 40 (1995) l. 4. O.C. Fagain andR. O'Kermedy, Biotech. Adv., 9 (1991) 351. 5. L.Gianfreda and M.R. Scarfi, Enzyme Stabilization:State of Art, Kluwer Academic Publishers, Netherlands, 1991. 6. J.H. Tae, Method. Enzymol., 91 (1983) 580. 7. G.E. Means and R.E. Feeney, Chemical Modification of Proteins, Holden Day Inc., California, 1971. 8. N. Nelson, J Biol. Chem., 153 (1944) 375. 9. Y.S. Rajput and M.N. Gupta, Enzyme Microb. Technol., 10 (1987) 143. 10. E.F. Hartree, Anal. Biochem., 48 (1972) 422.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
157
Studies on the stability of acid phosphatase (A. niger) by crosslinking with glutaraldehyde and soil humates N. Ortega, L. Berzal, M.D. Busto and M. Perez-Mateos Department of Biotechnology and Food Science, University of Burgos, Plaza Misael Bafiuelos s/n, Burgos 09001, Spain
1. INTRODUCTION Enzymes in the aqueous phase of soil are generally short-lived because they are easily inactivated by adsorption, denaturation or degradation [1]. Some enzymes, however, can persist for a long time because they are protected by humic colloids [2]. For example, urease [3], oxidases [4], proteases [5] or hydrolases [5] have been extracted from different soils as stable enzyme-humic complexes. In fact, we can regard humic matter as a highly charged polyelectrolyte having a random configuration and which is dominated chemically by a highly recalcitrant phenolic 'backbone' [7]. On the other hand, chemical cross-linking of enzymes can be easily achieved with bifunctional reagents (e.g. glutaraldehyde, which contain two identical reactive groups reacting with nucleophilic side chains of amino acids [8]). In this sense, Busto et al proposed to use glutaraldehyde to introduce functional groups into soil humic polymers which could then react covalently with water-soluble enzymes [9]. One part of the reagent is used to introduce an appropriate functionality into the reactive functional group from the polymer's backbone, and the other part is used to bind the enzyme. In the present work, the stability towards thermal and proteolytic deactivation of acid phosphatase native and immobilised by cross-linking with glutaraldehyde and soil humates is compared.
2. METHODS 2.1. Soil
Samples of an Umbric Dystrochrept soil were collected from the surface (0-10 cm), airdried, sieved (<2 mm) and stored at the field moisture level in sealed plastic bags at 4~ prior to use. The methods of soil analysis and physical and chemical characteristics of the soil (pH 4.3; CEC (cation exchange capacity) 0.17 cmol kg~; organic C 62.6 g kg~; C/N 14.9; sand 17.5%; silt 48.6%; clay 13.3%) together with their enzymatic activities have been fully described elsewhere [ 10].
158
2.2. Cell growth and enzyme production Spores from ,4. niger (strain obtained from the Spanish Culture Collection, CECT) were produced on Czapek-Dox agar slants incubated for 7 days at 30~ In order to produce the enzyme, 1 ml of a spore suspension was inoculated into 250-ml Erlenmeyer flasks containing I00 ml mineral medium (0.5 g NaNO3, 1.0 g K2HPO4, 0.5 g MgSO4-7H20 and 0.01 g FeSO4-7H20 per litre) at pH 7.0. This medium was supplemented with 0.2% peptone and 0.4% glycerophosphate (disodium salt pentahydrate) as carbon source. Cultures were incubated statically for 9 days at 30~ The fungal culture were filtered through membranes with a pore size of 0.45 ktm and the resulting mycelium washed with distilled water (10 ml), resuspended in citrate buffer pH 7.0 (10 ml) and disrupted 5 times for 60 s at 90 W in a Branson $250 sonicator (frequency 20 kHz) at 4~ The sonicated suspension was centrifuged at 22,700 x g for 30 min and the resulting supernatant was collected to assay the endocellular acid phosphatase activity. 2.3. Humates extraction Humates were obtained after 18 h of reciprocating agitation at 1.05 g (60 rpm) of samples obtained from the soil describe above, with 0.01 M pyrophosphate in a 1:5 soil:solution ratio (w/v) at pH 7.0-7.3. The humic acids were recovered in solution after 30 min of centrifugation at 15,000 rpm at 4~ and 0.45 ktm filtration (Filter Millipore HVLP OMTO5 Durapore| with 0.45 ktm retention, Minitan System) [4]. 2.4. Immobilisation procedure The acid phosphatase immobilised by cross-linking with glutaraldehyde and soil humates was prepared by mixing 3 ml of enzyme solution, 3 ml of humates solution extracted from soil and 4 ml of 2.5% glutaraldehyde solution (dissolved in 0.2 M Na-citrate buffer at pH 5.0). This reaction mixture was treated at 150 rpm in a rotatory shaker for 15 min at 45~ Final concentration of glutaraldehyde in the process of immobilisation was 1% [ 11]. 2.5. Phosphatase assay The activity of the soluble and immobilized acid phosphatase preparations were determined using p-nitrophenylphosphate (pNPP) as an artificial substrate. A reaction mixture containing 1 ml of native or immobilised enzyme, 3 ml of 0.2 M Na-citrate buffer (at pH 5.0) and 1 ml of 25 mM pNPP solution was incubated at 37~ for 1 h. The enzymatic reaction was stopped by adding 5 ml of 0.5 M NaOH and the organic matter was flocculated by adding 5 ml 0.5 M CaCl2. After filtration, the concentration of the p-nitrophenol (pNP) released was measured at 4 l0 nm using a spectrophotometer. Values shown in figures and tables represent the average of at least two triplicate assays. Duplicate controls to which substrate was added after incubation were assayed in all cases to deduct any non-enzymatic activity.
3. RESULTS AND DISCUSSION
3.1. Thermal stability The behaviour of the immobilised and free acid phosphatase against temperature is shown in Figure 1. Both enzymes had their highest activities at a temperature of 90~ Nevertheless, the
159 native enzyme showed an activity-temperature profile broader than that of the immobilised enzyme. This behaviour may be caused by the microenvironment effect of the humate matrix. Thermal stability of the immobilised and native enzyme was investigated by heating the enzyme for 1 h at several temperatures ranging between 30 and 90~ and the residual activity was measured by the standard assay method (Figure 2). Results showed that the free enzyme lost rapidly its activity when was incubated at temperature above 600C. In contrast, the phosphatase-humate complexes retained 25% of their activity after incubation for 1 h at 90~ This higher thermal activity could be due to a restriction on its degree of freedom caused by the immobilisation itself, consequently hindering heat denaturation [ 12].
1oo
.
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ua m
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~.0
4'0 6'0 8'0 TEMPERATURE,~
100 '~
Figure 1. Effect of the temperature on the free and immobilised acid phosphatase.
020
~klmmobilised
4'0 6C) 8'0 TEMPERATURE, ~
1()0 "~
Figure 2. Thermal stability of free and immobilised acid phosphatase.
Additionaly thermal stability of the free and immobilised enzyme at 60, 65, 70~ was also studied. This was determined by incubating the enzyme solution at these temperatures and withdrawing samples for assay at fixed intervals over an incubation period of 7 h. Thermal decay constants were calculated from a linear regresion analysis of semilogarithmic plots of the percentage activity remaining versus time. The half-lives of the native and immobilised acid phosphatase were calculated from the value of thermal decay constans (Table 1) [13]. The stabilisation factor (relationship between the half-live of immobilisation form and the native enzyme) at 60, 65 and 70~ for acid phosphatase-humic complexes were 4.8, 2.0 and 1.7, respectively.
3.2. Storage stability The storage stability is an important factor to be considered in applications of immobilised enzymes. In this work, the residual activity of free and immobilised enzyme was assayed after storage for 92 days at 4~ and -26~ (Table 2). It was found that the remaining activity of acid phosphatase from A. niger was 22% of the initial activity after a month of storage at 4~ and
160 Table 1 Half-lives of native and immobilised acid phosphatase Temperature t~a ~ Siabilisation (~ (h) factor b Enzy me 60 59.6 1.0 Native 65 13.2 1.0 70 2.6 1.0 60 288.3 4.8 Immobilised 65 26.7 2.0 70 4.5 1.7 " Apparent half-life (time required for acid phosphatase decline to 50% of its initial value). b Stabilisation factor is defined as the relationship between the half-lives of immobilised forms and of native enzyme. no activity was detected after two months. However, only 13% of the acitivity of the acid phosphatase-humic complexes crosslinking with glutaraldehyde was lost alter three months at 4~ On the other hand, the storage stability of the enzymes at -26~ was similar.
Table 2. . Storage stability of free and immobil!sed acid phosphatase at 4 and -26~ Storage time , Residual acid phosphatase (%) (days) Free Immobilised 4~C " _26oc 4~ '" _260(2 0 100.0 100.0 100.0 100.0 1 106.9 105.3 101.3 91.3 3 93.3 90.2 92.6 91.1 5 82.2 76.9 95.5 79.0 8 85.2 75.3 90.3 68.7 15 67.3 69.2 78.6 62.4 30 22.3 76.5 88.2 n.d. a 60 0.0 79.7 95.7 73.0 92 0.0 62.1 87.2 74.7 not determined. ,,
,,,
,
i
,
ii
,,
,i,
3.3. Stability towards proteolytic deactivation In the enzyme reactions that use crude enzyme preparations, inactivation of the enzyme is often accelerated by contamination with proteolytic enzymes. For instance, immobilisation is becoming increasingly recognized as a method of protecting biocatalysts from inactivation by proteolytic enzymes [14]. Pronase from S. griseus (0.5 mg ml1) was used for testing the resistance of the free and immobilised enzymes to proteolysis [ 15]. The results of Figure 3 show that the resistance to proteolytic deactivation of immobilised enzyme increased in comparison with the native enzyme. In fact, about 98% of the activity of free acid phosphatase was lost after 1 h exposure to protease, while the immobilised complexes
161 synthetised conserved 10~ of its activity after 24 h. In general, enzymes immobilised on synthetic supports are protected from proteolytic attack but to varying degrees depending on their mode of attachment [ 16]. IOO 8o
40
<:
20
ised ---.
Ol
o
;
Free
1'o
1'5 TIME,h
2'o
15 9
Figure 3. Effect of the pronase on the free and immobilised acid phosphatase.
REFERENCES 1. M. Perez-Mateos, M.D. Busto and C. Rad, J. Sci. Food Agric., 55 (1991) 229. 2. P.M. Huang and M. Schnitzed (eds.), Interaccion of Soil Minerals with Natural Organics and Microbes, Soil Science of America, Madison, 1986. 3. R.G. Burns, A.M. Pukite and A.D. McLaren, Soil Sci. Soc. Am. Proc., 36 (1972) 308. 4. M. Perez-Mateos, S. Gonzalez and M.D. Busto, Soil Sci. Soc. Am. J., 52 (1988) 408. 5. J. Mayaudon, L. Batistic and J.M. Sarkar, Soil Biol. Biochem., 7 (1975) 281. 6. M.D. Busto and M. Perez-Mateos, Biol. Fert. Soils, 20 (1995) 77. 7. R.G. Burns, Ann. Edafol. Agrobiol., 46 (1987) 1247. 8. S.S. Wong and L.J.-C Wong, Enzyme Microb. Technol., 14 (1992) 866. 9. M.D. Busto, N. Ortega and M. Perez-Mateos, Biores. Technol., 60 (1997) 27. 10. M. Perez-Mateos and S. Gonzalez, Biol. Fertil. Soils, 1 (1985) 153. l l.N, Ortega, M.D. Busto and M. Perez-Mateos, Abst. XX Congress SEBBM, Madrid (1997) 145. 12. I. Alkorta, C. Garbisu, M.J. Llama and J. Serra, Enzyme Microb. Technol., 18 (1996) 141. 13. S.K. Srivastava, K.S. Gopalkrishnan and K.B. Ramachandran, EnzymeMicrob. Technol., 6 (1984) 508. 14. L. Gianfreda, M. Modafferi and G. Greco, Enzyme Microb. Technol., 7 (1985) 78. 15. J.M. Sarkar and R.G. Burns, Soil Biol. Biochem., 16 (1984) 619. 16. O. Zaborsky (ed.), Immobilized Enzymes, CRC Press, Cleveland, 1973.
a This Page Intentionally Left Blank
Non-covalem processes in solution
a This Page Intentionally Left Blank
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Folding and association
versus
165
misfolding and aggregation of proteins
R. jaeni cke Institut for Biophysik und Physikalische Biochemie, Universit~t Regensburg, D-93040 Regensburg, Germany The acquisition of the spatial structure of proteins follows a "hierarchical condensation" mechanism, with local interactions between next neighbors and short-range weak interactions between domains and subunits accumulating to the marginal free energy generally observed for the functional state of globular proteins. Domains represent independent folding units such that the overall folding kinetics divide into the sequential collapse of subdomains and domains and their merging to form the compact t e r t i a r y structure. In proceeding to oligomeric proteins, docking of subunits follows the formation of "structured monomers". Thus, the self-organiza.tion of multimeric proteins may be quantitatively described by consecutive uni-bimolecular kinetics. Beyond a limiting protein concentration, aggregation (misassembly) will outrun proper domainpairing and subunit-association. In the cell, accessory proteins are involved in regulating the rate and yield of folding and association. Their technical application is expected to have a high impact on the downstream processing of pharmaceutically important proteins; related upscaling strategies gained from in v i t r o experiments are low-molecular weight additives, pulse-dilution and immobilization on matrices.
1. INTRODUCTION The topic of this article has three facets: the classical one referring to Max Perutz' description of the f i r s t in v i t r o renaturation experiments in terms of "unboiling the egg" [1], a second one. asking what folding experiments teach us about the self-organization of proteins in the cell, and a third one, aiming at biotechnological applications. In connection with the f i r s t problem, recent experimental developments give access to the whole range of processes from diffusion-controlled reactions in the nanosecond time range to slow isomerization and activation steps involved in molecular assembly or morphogenesis [2]. To the in v i t r o / in vivo issue, physical biochemistry cannot contribute much, except for model studies, (i) trying to mimic the cellular conditions, and ( i i ) explaining the molecular mechanisms of chaperone
166
action and folding catalysis [3.4]. At this point, not only cross-linking in s i t u and complementation in reticulocyte lysate come to mind, but also the in v i t r o translation of truncated mRNAand otherwise mutated mRNA[5]. Regarding the third issue, we have been witnessing that what started in the ivory tower 50 years ago has changed into industrial plants where cubic-meters of concentrated guanidine solutions are pumped in automated processes to yield grams of pharmaceuticals [6]. In the following review, all three aspects will be briefly discussed. 2. HIERARCHIESOF FOLDINGAND STABILITY
In the structural hierarchy of proteins, the different levels refer to both folding and stability. Increasing packing density, by improved van der Waals interactions and water release from hydrophobic residues, provides enthalpic and entropic increments of the free energy of stabilization AGu_~N. This represents only a marginal difference between the attractive and repulsive forces characteristic for the s t a b i l i t y of proteins in their native state. Considering the numbers involved, 5000 atoms making up an average protein molecule give a AGu_~N value of no more than -60 kJ/Mol, i.e., the equivalent of just a few weak interactions [7]. The biological significance of this observation is threefold: (i) optimization of the structure-function relationship in the course of evolution is aimed at f l e x i b i l i t y (catalysis, regulation, turnover) rather than s t a b i l i t y ; ( i i ) under physiological conditions, native globular proteins are generally at the borderline of denaturation, ( i i i ) since the native state is a state of minimum potential energy, folding intermediates must exhibit even lower s t a b i l i t y than their native counterparts, so that misfolding and subsequent kinetic competition of reshuffling and off-pathway reactions are expected to Occur. Evidence that protein biosynthesis and folding in the cell do not yield 100 % came from turnover experiments. For example, the tailspike protein of bacteriophage P22, even under optimum growth conditions of its host yields less than 50%; under unbalanced physiological conditions, only wrong conformers are produced which are continuously removed by proteolysis. At this point, chaperones come into the play. In correlating the i n t r i n s i c s t a b i l i t y with the structural hierarchy, thermodynamic measurements on point mutants, protein fragments and homologs differing in their state of association have clearly shown that each structural level makes its own contribution [7]. As proven by NMR and other spectroscopic techniques, oligopeptides may form stable non-random conformations; at
167
a minimum length of 15 residues, they have been shown to sustain native-like structure. Based on this observation, short helices are considered to play a role as 'nucleating' elements in the folding process. Larger fragments such as domains are known to 'fold-by-parts' [10]. Even at the level of subdomains, they may exhibit intrinsic s t a b i l i t i e s not far from the free energies observed for the uncleaved parent molecules [8]. Further stabilization has been found at the subunit level. For domains and subunits, yB-crystallin and lactate dehydrogenase (LDH) may serve as examples (Figure 1). In the case of LDHo the s t a b i l i t y decreases steadily, from the highly stable (native) tetramer down to domain fragments The 'proteolytic dimer' requires structure-making salts to exhibit a c t i v i t y , whereas the monomer is inactive and only detectable as a short-lived folding intermediate on the pathway of reconstitution" the separate NAD- and substrate-binding domains are unstable, but s t i l l s u f f i c i e n t l y structured to recognize each other, exhibiting a mutual 'chaperone effect' during j o i n t reconstitution (Figure 1E) [11].
g 0t ,o .
'~ O:
o
1 ~' " ~C ,oo
o
I
!
J
Oo ~
9
~~ 9
OL.,.,.,.,.,.,-,. 0 2 L,
9
6
c u (M)
0
g
I 2 3 CGdmCI {M)
0
50 100 150 Time {rain)
8 C
Figure 1. Denaturation-renaturation of domain- and subunit-proteins. A: Ureainduced equilibrium transitions of monomeric yB-crystallin (o) and its monomeric linker mutant, with the bent yB-peptide between the domains replaced by the extended l i n k e r of the 'domain-swapped', dimeric BB2-crystallin (e), showing that domain interactions dominate over subunit interactions. B: Denaturation of the separate N- (o) and C-terminal (e) domains, i l l u s t r a t i n g mutual domain stabilization. C: Domain swapping of monomeric yB-crystallin (upper two domain, fat line) causes dimerization of BB2-crystallin (thin line) [3,8]. D: Denaturation of tetrameric (e) and (nicked) dimeric LDH (o). E: Reactivation of p r o t e o l y t i c dimers (El) and equimolar amounts of the 14 and 21 kDa domain fragments after denaturation in 6 M GdmCl; separate (o) and j o i n t reconstitution (e) [11].
168
Extrinsic factors such as ions, cofactors and non-proteinaceous components (e.g.,nucleic acids or carbohydrates) may contribute significantly to protein s t a b i l i t y ; they may also be of importance in determining the mechanism of folding and the state of association [3]. 3. MECHANISMOF FOLDINGAND ASSOCIATION
The elucidation of the folding mechanism of a protein is an insoluble problem, because i t requires the complete description of the nascent (unfolded) and final (native) states, together with all intermediates along the U ~ N transition. The main obstacle in reaching this goal is the elusive nature of the folding protein. The fact that protein folding must occur within a biologically feasible time span, i.e. a time much shorter than the life-time of an organism, clearly excludes a random search mechanism over all conformational space, forcing us to assume that there must be kinetic pathways of folding. Whether they follow a 'framework model', according to U ---~---~ I i
~ N
(1)
or multiple pathways has become a controversial issue after Monte Carlo calculations resulted in 'folding funnels' instead of sequential reactions, indicating that the I i ensemble represents off-pathway dead-end traps rather than true intermediates. Yet, from a large body of experimental data, i t seems clear that the l i ' s a r e significantly populated, but not necessarily synchronized [12], supporting the idea that evolution has selected proteins for robust intermediates, this way allowing efficient folding [13]. Advances in spectroscopic methods laid the fundament to a generalized folding mechanism. This involves as a f i r s t step next-neighbor interactions at nucleation sites, followed by a hydrophobic collapse to a state containing native-like secondary structure, but s t i l l non-native t e r t i a r y interactions. Shuffling of this 'molten globule' leads to the correct tertiary fold, either at the domain or subunit level. The final step is then the minimization of the remaining hydrophobic surface either by domain or subunit interactions. The latter requires the formation of the correct subunit interfaces in the 'structured monomers' in order to guarantee specific pairing. At this point, the occurrence of domains as independently folding entities may provide the solution of one of the challenges evolution has been facing in proceeding from small proteins to large protein assemblies, namely the formation of tight van
169
der Waals interactions between two polypeptide chains. Here, 'domain swapping' a f t e r gene d u p l i c a t i o n offers a mechanism which allows at least homooligomeric assemblies to be easily explained [14]. Figure 1C illustrates the idea, using the By-crystallin family as an example. Small proteins or domains as constituents of large proteins commonly collapse In a highly cooperative manner into a compact structure. This is nativelike, although there have been reports showing that non-native intermediates may occur [15]. As mentioned, fast secondary structure formation precedes slow multi-step rearrangements observable on the seconds to minutes time scale [16]. Due to limited time resolution and the multi-step and multiple pathway problems, a complete analysis of the folding kinetics has not been achieved for any protein so far. The most detailed mechanisms presently available refer to small single-chain one-domain proteins [17]. General conclusions from these model systems are the following: ( i ) in agreement with the above two-phase mechanism, there are compulsory pathways of folding which are, at least in part, sequential; (i i) secondary structure formation is driven by local hydrophobic surface minimization and precedes t e r t i a r y structure formation, and (i i i) t e r t i a r y interactions become increasingly defined as water release consolidates the hydrophobic core. In proceeding to domain proteins and protein assemblies, the previous conclusions remain widely unchanged. The reason is that proteins 'fold by parts', i . e . , domains fold and unfold independently according to [Ni-Nj]
~
Ni-Uj
~Ui-U
j
(2)
where i and j refer to different domains in their native (N) and unfolded (U) states (Figure 1A). Obviously, this observation holds not only in v i t r o but also in the cell [8]. The mechanism is most significant, not only in connection with the protection of the nascent polypeptide chain from proteases and the evolution of multi functional enzymes, but also as a contribution to the rate enhancement of protein folding. There are examples where biological function requires the cooperation of domains, e.g.. two domains forming one active center. In such cases, domain pairing [Ni ] - [ N j ] ~
[Ni-Nj]
(3)
may occur as an additional rate-limiting step in the overall folding reaction.
170
In oligomeric proteins, subunit assembly corresponds to domain pairing. The preceding steps on the pathway are those described before, with structured monomers (with complementary subunit interfaces) as the final product before association can take place. Evidently, their c o l l i s i o n complex may undergo intramolecular rearrangements to reach the state of maximum packing density and minimum hydrophobic surface area. Thus, for a dimer, folding and association may be a sequential uni-bi-unimolecular reaction, according to fast 2J~,
kI
;
2 M'
k2 -
2 M~
k1 M2
>
(4)
N
with J(, M', M as unfolded, intermediate and structured monomer, N as native dimer and k1, k2 as I st- and 2nd-order rate constants. How the intermediates along the pathway can be monitored depends on the specific structure-function relationship for a given quaternary structure. In most cases, biological function requires the native state of association so that the final ratedetermining step can be monitored by measuring activity. Preceding steps may be accessible to spectral analysis, H-D exchange, cross-linking, HPLC and a wealth of other methods [18,19]. Figure 2 i l l u s t r a t e s the reactivation of cytosolic (s-MDH) and mitochondrial malate dehydrogenase (m-MDH), showing that closely related proteins may obey different folding-association mechanisms: in the case of s-MDH, dimerization is diffusion controlled, whereas m-MDH follows sequential uni-bimolecular kinetics [3]. 100 i'.-.,.b. A
n,-
0 0
.
-1 1~176B;.,-,oH
=f- - .._~f. - ' - ~ "--..
loo i -
. =
~
I < "':' i Oo "[/=I
I , , , 0 , I 9 ,-0.5 1 1.5 2-2/, 0 0.5 1 1.5 2 6 1 0 0--0.5 1 1.5 2 Time (h) Time (h) Time (h) Figure 2. Reactivation kinetics of porcine s-MDH and m-MDH. A. s-MDH: Denaturation in 6 M GdmCl renaturation in phosphate buffer pH 8 at D~ 3 40-300 nM shows no concentration dependence. B. m-MDH: Denaturation at . or in 5 M GdmCl: renaturation as in A: CMnU= 2 (I). 4 ( D ) , 10 (Z~), 35 (0), 88 ( = ) and 143 nM (A). Curves c a l c u l a ~ ] according to a uni-bimolecular mechanism. C. m-MDH. Reactivation at CMDH,= 65 nM after denaturation at pH 2 (o), in 6 M GdmCl (m). 8 M urea ( A ) . anG in the presence of 10 mM NAD+ (>95% saturation) ( 0 ) [1,3].
171
4. OFF PATHWAYREACTIONS
There are three stages where side reactions on the folding pathway may compete with proper folding and association the hydrophobic collapse, the merging of domains and the docking of subunits. Examples are- ( i ) transient interactions in the collapse of cytochrome c or B-lactoglobulin, ( i i ) domain mismatch of tryptophan synthase B2 and octopine dehydrogenase, and (i i i) inclusion body formation. At all three levels, 'recognition' is involved in the sense that specific substructures or surfaces have to be pre-formed in order to proceed on the folding path toward maximum packing density and minimum hydrophobic surface area. Collapse and domain merging involve intramolecular rearrangements. Due to the high local concentration of the reacting groups or surfaces within one and the same polypeptide chain, they are not significantly affected by neighboring molecules, i . e . , they obey I st - order kinetics with the slowest isomerization reaction determining the overall rate. In the case of domain proteins, the r e l a t i v e s t a b i l i t i e s of the domains and the contributions of the domain interactions to the overall s t a b i l i t y are crucial. The significance of the linker peptide connecting two well-defined domains has been studied by "grafting" experiments, e.g., by mutually exchanging the linker peptides of B- and v-crystal l in (Figure 1A) [8]. In both transplants, domain contacts dominate over subunit contacts. The recombinant separate domains do not interact with each other, stressing the local-concentration argument. Evidently, separate domains or other substructures may very well recognize each other and form complexes' RNaseS [20] and the folding and association of nicked lactate dehydrogenase [11] may serve as examples. The fact that in the l a t t e r case the yield does not exceed 15 % illustrates the importance of side reactions. In the case of the related monomeric octopine dehydrogenase, kinetic partitioning upon reactivation yields 70% native, and 30 % non-native protein [21] ~ U - ~- I N ~ IN
N
(70%)
(5)
(30%)
It is assumed that I N is "irreversibly denatured" as a consequence of wrong domain interactions. Repetitive denaturation/renaturation yields 70% N per cycle so that after a 2nd and 3rd cycle only ca. 50 and 35% of the original a c t i v i t y are recovered.
172
Obviously, i t is the kinetic partitioning between folding/association on one side, and misfolding/misassembly, on the other that determines the wellknown decrease in the yield of reactivation at high protein concentration. In going from single-chain domain proteins to protein assemblies, i t is wellestablished that this side reaction becomes the dominant effect in the overall mechanism of folding and association (Figure 3A) [22]. The underlying kinetic mechanism for an oligomer made up of n subunits is:
(m+n)~--~
kI k2 (m+n) M'- > n M ~ Mn,native (6)
v({ m
M'm
Here, ~6~, M', and M stand for the unfolded, collapsed and structured monomer, k1, k2 and k>2 for r a t e - l i m i t i n g I st-, 2nd and higher than 2nd-order rate constants, and ~m and M m for the products of aggregation as side reaction of folding/association. Evidently, the latter equation corresponds to Equ. (4) which describes the limiting case at high dilution at which premature c o l l i s i ons of misfolded molecules can be ignored, as shown in Figure 3B, i t is sufficient to quantitatively describe the observed inverse concentration dependence of reactivation and aggregation. Considering the underlying molecular mechanism, three questions need to be answered. ( i ) what is the committed step in aggregate formation, i . e . , at which stage along the sequential reaction are aggregates formed, ( i i ) when is the structured monomer committed to end up as the native protein and (i i i) what is known about the structure of aggregates and their constituent polypeptide chains. With respect to the f i r s t two problems, commitment to aggregation was shown to be a fast reaction, whereas the kinetics of the 'commitment to renaturation' followed precisely the slow kinetics of the overall reactivation. This means that there are fast precursor reactions on the folding path (collapsed states) all of which allow s t i l l aggregation, whereas. after a certain intermediate has been formed, slow shuffling leads 'one way' to the native state [23]. Regarding the third question, i . e . , the structure of aggregates, electron microscopy and circular dichroism indicate that wrong subunit interactions give rise to irregular networks with a broad distribution of highly structured particles at least 10 times the size of the native proteins. They resemble the native protein in its spectral properties, as far
173
as turbidity allows this conclusion (Figure 3A). For structural characteris tlcs of inclusion bodies and protein deposits, cf. [22,24]. A
B
.. I00~ ~ "~ 75
C
.
I
100 .,.,.
so
0
001
01
1.0
ClD H (pM)
8--_1
10
_--
01
5O
~
0001 001
~
0.1
O
10
~
DO lpM)
10
113(3
0
1.0
L- Arginine (M]
20
Figure 3. Kinetic partitioning between folding and aggregation. A. % reactivation (o), renaturation (Q) and light scattering (Z~) for porcine muscle LDH; denaturation at pH 2 renaturation at pH 7 (10~ [3,17] B. Reactivation calculated according to the competition of lSZ-order folding and diffusioncontrolled aggregation [16]. C. Effect of arginine on the reactivation and reoxidation of tissue plasminogen activator [25].
5. KINETICPARTITIONING. PRACTICALAND CELLULARASPECTS 5.1. Aggregation and inclusion body formation As has been mentioned, aggregation in vitro and inclusion body formation in the cell correspond to each other 9overexpression leads to high local concentrations of folding, yielding precipitates instead of native protein. One approach to cope with this problem is the use of weaker promotors, this way reducing the concentration. Since the aggregation reaction is of >2nd order, this may lower the local level of aggregation-competent folding intermediates below a c r i t i c a l concentration, thus favoring correct folding and association. However, there are two reasons why this approach is of no practical use- f i r s t , the overall yield of the recombinant protein per gram cell mass is drastically decreased, and second, its purification requires a f u l l scale separation of the guest molecule from the bulk of the host proteins, whereas inclusion bodies with their characteristic low heterogeneity allow highly simplified purification procedures for their fractionation. Thus, in many cases, down-stream processing starting from inclusion bodies has been the method of choice. In order to work at sufficiently low protein concentration during reconstitution, discontinuous 'pulse dilution' may be applied a certain amount of the protein is subjected to dilution and reactivated at the
174
sub-critical protein concentration: approaching the final value of reconstitution, the next portion of the concentrated solution of the denatured protein is added, and so on until the whole batch is transferred. Apart from keeping the concentration of the denatured protein low, this approach has the advantage that the increasing level of the renatured protein exerts a stabilizing effect on folding intermediates comparable to the one used routinely by adding, e.g.. serumalbumin. In this context, additives such as arginine may strongly increase the yield by shuffling aggregates back on the productive folding path (Figure 3C). L i t t l e is known about specific groups involved in the aggregation reaction. Early systematic experiments suggested hydrophobic interactions as well as covalent disulfide linkages to be of major importance. Recent in v i t r o and in vivo studies confirm this result [22,25,26]; test cases have been rhodanese and bovine growth hormone: for the f i r s t , detergents were shown to compete with aggregation, whereas BGH mutants with enhanced hydrophobic surface area showed enhanced aggregation [22,24-28]. The partitioning between folding and aggregation has been most intensively studied in the case of the tailspike protein (Tsp) from Salmonella bacteriophage P22 and the numerous mutants of this protein which either increase or suppress aggregation ( Figure 4) [30]. The wildtype trimer is highly stable and shows a close similarity in its in vivo and in v i t r ~ folding behavior. Upon release from the ribosome or upon dilution from denaturant solutions, A
B
C
~ams
Figure 4. Structure and folding mechanism of Tsp from bacteriophage P22. A. Polypeptide backbone of the truncated tailspike trimer, one subunit traced in bold. The molecule is folded into a right-handed parallel #-helix. B shows a section through the #-helix part, illustrating the side-by-side association of the subunits. C. Scheme for the in v i t r o and in vivo assembly of Tsp [22.29].
175
the polypeptides fold into an assembly-competent conformation bulk of the #sheet secondary structure and their aromatic amino-acid side chains are close to the native state, but even as 'protrimers' they are s t i l l highly unstable and prone to aggregation. The anomalous s t a b i l i t y is only acquired in a slow rearrangement reaction when the intertwined parallel #-helices merge to form the native trimer. Correspondingly. a significant part of the Tsp folding reaction occurs after subunit association. During maturation in vivo as well as refolding in v i t r o , the fraction of chains capable of maturing to native Tsp decreases with increasing temperature, the rest aggregates. Temperaturesensitive folding ( t s f ) mutations reduce the folding yield at elevated temperatures, whereas second-site suppressor mutations (su) improve folding under such conditions. Both types of mutations act by altering the s t a b i l i t y of tailspike folding intermediates [22,29].
5.2. Reconstitution in the presence of accessory proteins Inclusion bodies are the product of intracellular aggregation. They differ from aggregates formed in the test tube by their high packing density and large size which may sometimes span the entire diameter of the cell [23]. As a consequence, they can be easily harvested and washed by fractionated centrifugation, favoring their use in down-stream processing of recombinant proteins. The isolation of the desired protein follows exactly the same routine of in v i t r o denaturation-renaturation described before. For detailed guidelines, cf. [17,26]. Recently, the repertoire of methods has been extended by attempts to mimic in vivo conditions with respect to folding catalysts and chaperone proteins. Three examples may serve to illustrate the results. The f i r s t refers to the GroE system, the major molecular chaperone from E. coli and, at the same time, the role model of chaperone action. The molecule represents a K+- dependent weak ATPase, consisting of GroEL, a double-disc with 7-fold symmetry and 14 identical 60 kDa subunits, forming a hollow cylinder, which is closed by one or two heptameric GroES-lids serving as ATPase inhibitors. ATP. ADP and the substrate-protein are involved in drastic conformational changes that alter the interior of the cage from strongly hydrophobic to hydrophilic. As long as stretches of hydrophobic residues of the folding polypeptide chain are exposed, this shows high a f f i n i t y to the chaperone. During the off-phases, 'annealing' continues until the substrate has reached its native state. This does not bind to the chaperone. Thus. considering the kinetic partitioning between folding and aggregation, the chaperone does nothing but keeping aggre-
176
gation-competent folding intermediates below a c r i t i c a l concentration [30]. This simple explanation is confirmed by in v i t r o experiments using c i t r a t e synthase (CS) as substrate [31]. After unfolding and subsequent d i l u t i o n of the denaturant, only a small portion of the enzyme regains activity, while the major part aggregates. Renaturation in the presence of GroEL leads to the formation of a stable binary complex with non-native CS. thus i n h i b i t i n g reactivation completely. Only after GroES and ATP are added, the enzyme is released and reactivated. Stoichiometric amounts of GroEL are s u f f i c i e n t to block aggregation. The slow increase in light scattering after the addition of GroES and ATP shows that the released protein is s t i l l in its non-native state, prone to unspecific aggregation (Figure 5). The previous experiments clearly show that the GroE system does not rescue aggregated protein, nor is i t a folding catalyst accelerating the rate of folding. On the contrary, in the case of CS, reactivation is slowed down, probably due to the binding and release processes taking place during i t e r a t i v e annealing. For a detailed discussion of chaperones and their various modes of action, see [22,30,32.33]. The second example refers to the reactivation of a denatured and reduced immunotoxin (B3(Fo)-PE38KDEL) composed of the VH region of a carcinomaspecific antibody, connected by a f l e x i b l e linker to the corresponding VL chain, which is in turn fused to truncated Pseudomonas exotoxin. The chimeric protein contains three disulfide bonds, one in each antibody domain, and one in the toxin part. Upon renaturation, aggregation of non-native polypeptide A 9
c75
9- - -
.9
o
o:
B 75~
~
=
....
._c
._o
.,.
.--
75
~,
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25
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0
' ~' 0 100 200 300 400 Citrate s y n t h a s e ( n M )
"
~
C 60
._
; ~
o
~ ,
~ 20
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i 0
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2 Time
i /, 6 (mJn)
Figure 5. Reconstitution of porcine citrate synthase after preceding denaturation in 6M GdmCl. A. Concentration dependence of reactivation (o) and aggregation (e). B. Reactivation in the presence of GroEL without GroES and ATP ( B ) , with GroES and ATP (e), and with the addition of GroES and ATP after 30 (~7) and 60 min ( w ) , respectively. C. Aggregation of the enzyme during reconstitution in the absence of GroE (o), in the presence of GroEL ( n ) , and in the presence of GroEL/GroES and ATP; addition at zero time of reconstitution ( w ) . after 30 sec (e), and after 4 min ( v ) [31].
177
chains and the formation of incorrect disulfide linkages lead to >90 % inacrive molecules. In order to improve the yield, E. Coli GroE and DnaK as well as protein disulfide isomerase (for SH/SS exchange) were applied. Both GroE and DnaK are found to influence the reaction: GroEL alone inhibits reactivation, whereas the complete GroEL/ES system significantly increases the yield of active protein. DnaK exhibits the same effect. PDI synergistically stimulates the reactivation, duplicating the yield compared to the non-enzymatic disulfide bond formation [34]. The third example deals with the renaturation, purification and down-stream processing of recombinant antibody fragments expressed in E.Coli. In this case, temperature, protein concentration, redox buffers, folding catalysts and destabilizing additives were varied in order to optimize the renaturation process [35,36]. It turned out that, in the case of the oxidized Fab fragment, peptidyl prolyl isomerases accelerate the refolding reaction and, at the same time, increase the yield of the native final product, thus proving that catalysis affects the kinetic partitioning by enhancing a rate-limiting step on the folding path. 5.3. Immobilization of proteins Folding/unfolding of immobilized proteins not only allow to avoid the off-pathway aggregation reaction but also opens the way to study the vectorial character of protein self-organization by comparing the results of folding starting from the N- or the C-terminal end of the polypeptide chain. Using ~glucosidase with polyionic N- or C-terminal t a i l s to reversibly immobilize the protein to an ion-exchange resin as a model, i t was shown that the enzyme (with an Arg 6 t a i l ) shows long-term s t a b i l i t y over weeks. Its reactivation yield, after preceding guanidine denaturation, is increased at least 5-fold, and the upper l i m i t of protein concentration without aggregation is shifted from-10 ~g/ml to -5 mg/ml [37]. The fact that both refolding from the C- and N-terminal end yield active enzyme is clear evidence that the vectorial character of protein biosynthesis, i . e . , the cotranslational folding from the Nto the C-terminal end of the polypeptide chain, does not determine the threedimensional structure of proteins. 5.4. Cel I ul ar aspects As taken from the classical examples of protein folding studies, three steps may be r a t e - l i m i t i n g in the self-organization of proteins, disulfide shuffling, proline cis-trans isomerization and association. On the other hand.
178
chains and the formation of incorrect disulfide linkages lead to >90 % inactive molecules. In order to improve the yield. E. Coli GroE and DnaK as well as protein disulfide isomerase (for SH/SS exchange) were applied. Both GrcE and DnaK are found to influence the reaction: GroEL alone inhibits reactivation, whereas the complete GroEL/ES system significantly increases the yield of active protein. DnaK exhibits the same effect. PDI synergistically stimulates the reactivation, duplicating the yield compared to the non-enzymatic disulfide bond formation [34]. The third example deals with the renaturation, purification and down-stream processing of recombinant antibody fragments expressed in E.Coli. In this case, temperature, protein concentration, redox buffers, folding catalysts and destabilizing additives were varied in order to optimize the renaturation process [35.36]. It turned out that, in the case of the oxidized Fab fragment. peptidyl prolyl isomerases accelerate the refolding reaction and, at the same time, increase the yield of the native final product, thus proving that catalysis affects the kinetic partitioning by enhancing a rate-limiting step on the folding path. 5.3. Immobilization of proteins Folding/unfolding of immobilized proteins not only allow to avoid the off-pathway aggregation reaction but also opens the way to study the vectorial character of protein self-organization by comparing the results of folding starting from the N- or the C-terminal end of the polypeptide chain. Using ~glucosidase with polyionic N- or C-terminal tails to reversibly immobilize the protein to an ion-exchange resin as a model, i t was shown that the enzyme (with an Arg 6 t a i l ) shows long-term s t a b i l i t y over weeks. Its reactivation yield, after preceding guanidine denaturation, is increased at least 5-fold, and the upper l i m i t of protein concentration without aggregation is shifted from-10 pg/ml to -5 mg/ml [37]. The fact that both refolding from the C- and N-terminal end yield active enzyme is clear evidence that the vectorial character of protein biosynthesis, i . e . . the cotranslational folding from the Nto the C-terminal end of the polypeptide chain, does not determine the threedimensional structure of proteins. 5.4. Cel I ul ar aspects As taken from the classical examples of protein folding studies, three steps may be rate-limiting in the self-organization of proteins, disulfide shuffling, proline cis-trans isomerization and association, On the other hand.
179
there are two enzymes, localized in the appropriate cellular compartments to catalyze the f i r s t two steps, and there are 'machineries' of chaperones and 'cohortes' of helper proteins to assist the third reaction [16.30.32,33]. The development in all three areas during the last few years has been breathtaking. In spite of that, both folding catalysis and chaperone action are far from being understood. Only a few aspects w i l l be outlined in the following paragraphs. For details reference is made to a number of recent monographs and reviews [5,26,38,39]. The biological significance of protein disulfide isomerase (PDI), peptidyl prolyl cis-trans isomerase (PPI) and chaperones is now well-established, and structural data for representatives of all three types of accessory proteins have been reported with s u f f i c i e n t l y high resolution so that mechanistic information is available. DsbA, the PDI-homolog in Eocoli, shows a close relationship to thioredoxin. Its second domain may be involved in the communication with a more complex machinery, including a membrane-spanning unit (DsbB) involved in the regeneration of DsbA. The anomalous redox and s t a b i l i t y properties of DsbA (with the reduced form of the enzyme showing higher s t a b i l i t y than the oxidized one) have been elegantly resolved by mutant studies [40,41]. Apart from its PDI activity, the enzyme acts as a subunit in a number of other enzyme complexes the common denominator seems to be that in al I cases recognition of nascent polypeptide chains is involved. PPIs seem to be ubiquitous enzymes catalyzing the rotation around the X-pro peptide bond and inhibiting signal transduction processes. The three-dimensional structures of two different representatives of the PPI family have been elucidated, supporting a "catalysis-by-distortion" mechanism. The substrate specificity is low. Obviously, in the cell, PPIs serve various functionstheir function as trigger factor and their location in the ER, as well as their effect on the formation of collagen in fibroblasts support the assumption that they are involved in in vivo folding. Considering the complex regulatory processes in higher cells, definitive proof, e.g., in connection with the maturation of collagen, s t i l l needs further investigation [42-44]. With respect to chaperones, even for the best-known systems, DnaK/DnaJ/GrpE and GroEL/GroES, significant information is s t i l l lacking. Therefore, generalizations have to be taken with care. This holds (i) for the stoichiometry and role of co-chaperones and the components in the cycle of ATP hydrolysis, (i i) for the apparent lack of substrate speci f i c i t y . (i i i ) for the conformational state of the bound polypeptide, (iv) for the interaction with other cellular
180
components toward 'chaperone machines', (v) for the question whether chaperones catalyze protein folding, acting as helper proteins and "unfoldases", etc. Obviously, "resurrection" of misfolded and misassembled proteins ( i f i t occurs) is the exception rather than the rule, for steric reasons, because of the limited space in the "cage", and for energetic reasons, because aggregates are commonly trapped in a deep energy well. Regarding the state of association of the released protein substrate, direct proof for the monomeric state, i .e. against oligomerization on the chaperone, came from reconstitution experiments with glutamin synthetase, where the formation of the native dodecamer has been shown to occur in solution, after release of the subunits from the binary GroEL-complex [45]. What drives the release, and how the polypeptide chain in the chaperone-substrate complex looks like, is s t i l l unresolved [30]. In summary, the picture that emerges from studies on pro- and eukaryotes clearly shows that off-pathway reactions in the cell are blocked by a whole arsenal of components from the moment of translation. The reason why, in spite of this protection, protein biosynthesis and subsequent structure formation commonly does not yield 100% is obvious: interactions involved in chaperone action must not be too strong, because otherwise the nascent polypeptide chain would be trapped. Thus, during the off-reaction, molecules may escape from the folding path, ending up in degradation. There are a number of aspects of in vivo folding that have not been considered: genome organization, nucleic acid secondary structure, codon usage, cotranslational modification, etc. The reason is that at present there is no clear evidence proving any of these to play a significant role in the folding/association and misfolding/ misassembly reactions of proteins in the cellular environment. ACKNOWLEDGMENTS Work performed in the author's laboratory has been generously supported by Grants of the Deutsche Forschungsgemeinschaft, the Fonds der Chemischen Industrie, the Alexander von Humboldt Stiftung, the Max Planck Gesellschaft and the European Community.
REFERENCES 1. 2. 3. 4.
R. Jaenicke (ed.), Protein Folding, North Holland, Amsterdam, 1980. C.M. Dobson and O.B. Ptitsyn (eds.). Curr. Opin. Struct. Biol. 7 (1997) I. R. Jaenicke, Progr. Biophys. Mol. Biol. 49 (1987) 117. R. Jaenicke, Biol. Chem. 379 (1998) 237.
181
5. R.B. Freedman, in Protein Folding (T.E. Creighton, ed.), W.H. Freedman, New York, 1992. 6. R. Rudolph, in Protein Engineering: Principles and Practice (J.L. Cleland and C.S. Craik. eds.), 283, Wiley-Liss Inc., New York, 1996. 7. R. Jaenicke, Eur. J. Biochem. 202 (1991) 715. 8. R. Jaenicke, Progr. Biophys. Mol. Biol. (1998) in press. 9. R. Jaenicke, FASEB J. I0 (1996) 84. I0. D.B. Wetlaufer, Proc. Natl. Acad. Sci. USA 70 (1973) 697. i i . D.V. Laurents and R.L. Baldwin, Biophys. J. (1998) in press. 12. R.Lo Baldwin, J. Biomol. NMR 5 (1995) 103. 13. M.P. Schlunegger, M.J. Bennett and D. Eisenberg, Adv. Prot. Chem. 50 (1997) 61. 14. R. Jaenicke, Biochemistry 30 (1991) 3147. 15. T. Kiefhaber, R. Rudolph, H.-H. Kohler and J. Buchner, Bio/Technology 9 (1991) 825. 16. R. Jaenicke, Curr. Topics Cell. Reg. 34 (1996) 209. 17. R. Rudolph, G. BOhm, H. Lilie-and R. Jaenicke, in Protein Structure: A Practical Approach, 2nd Ed. (T.E. Creighton, ed.), 57, IRL, Oxford, 1997. 18. W.A. Eaton, V. Munoz, PoA. Thompson, C.-K. Chan and J. Hofrichter, Curt. Opin. Struct. Biol. 7 (1997) I0. 19. F.M. Richards and P.J. Vithayathil, J. Biol. Chem. 234 (1959) 1459. 20. U. Opitz, R. Rudolph, R. Jaenicke, L. Ericsson and H. Neurath, Biochemistry 26 (1987) 1399. 21. W. Teschner, R. Rudolph and J.-R. Garel, Biochemistry 26 (1987) 2791. 22. R. Jaenicke and R. Seckler, Adv. Protein Chem. 50 (1997) I. 23. M.E. Goldberg, R. Rudolph and R. Jaenicke, Biochemistry 30 (1991) 2790. 24. G. Georgiou, ACS Symp. Ser. 516 (1993) 53. 25. R. Rudolph, in Modern Methods in Protein and Nucleic Acid Research (H. Tschesche, ed.) 149, de Gruyter, Berlin, New York, 1990. 26. A. Mitraki and J. King, Bio/Technology 7 (1989)690. 27. S. Tandon and P.M. Horowitz, j . Biol. Chem. 261 (1986) 15615. 28. D.N. Brems, S.M. Plaisted, H.A. Havel and C.-S.C. Tomich, Proc. Natl. Acad. Sci. USA 85 (1988) 3367. 29. A. Mitraki and J. King, FEBS Lett. 307 (1992) 20. 30. M. Beissinger and J. Buchner, Biol. Chem. 379 (1998) in press. 31. J. Buchner, M. Schmidt, M. Fuchs, R. Jaenicke, R. Rudolph, F.X. Schmid and T. Kiefhaber, Biochemistry 30 (1991) 1586.
182
32. A.L. Fink and Y. Goto (eds.) Molecular Chaperones in the Life Cycle of Proteins, M. Dekker, New York, 1998. 33. B. Bukau (ed.) Molecular Biology of Chaperones, Harwood, Amsterdam. 1998. 34. J. Buchner, U. Brinkmann and I. Pastan. Bio/Technology I0 (1992) 682. 35. H. Lille, K. Lang, R. Rudolph and J. Buchner, Protein Sci. 2 (1993) 631. 36. H. Lilie, S. McLaughlin, R.B. Freedman and J. Buchner. J. Biol. Chem,269 (1994) 14290. 37. G.Stempfer, B. HOll-Neugebauer, E. Kopetzki and R. Rudolph, Nature Biotechnol. 14 (1996) 329, 481. 38. C.B. Anfinsen, J.T. Edsall, F.M. Richards and D.S. Eisenberg (eds.), Accessory Folding Proteins, Adv. Protein Chem. 44, Academic Press, San Di ego, 1993. 39. S. Betts, C. Haase-Pettingell and J. King, Adv. Protein Chem. 50 (1997) 243. 40. J.C.A. Bardwell, Mol. Microbiol. 14 (1994) 199. 41. M. Huber-Wunderlich and R. Glockshuber, Folding and Design, (1998) in press. 42. T. Zarnt, T. Tradler, G. Stoller, C. Scholz, F.X. Schmid and G. Fischer, J. Mol. Biol. 271 (1997) 827. 43. F.X. Schmid, L. Mayr, M. MOcke and E.R. SchOnbrunner, Adv. Protein Chem. 44 (1993) 25. 44. G. Fischer, T. Tradler and T. Zarnt, FEBS Lett. (1998) in press. 45. M.T. Fisher, J. Biol. Chem. 268 (1993) 13777.
Stability and Stabilizationof Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
183
Activity o f m o n o c l o n a l antibodies in prevention o f in vitro aggregation o f their antigens Beka Solomon, Tamar Katzav-Gozanski, Rela Koppel and Eilat Hanan-Aharon Department of Molecular Microbiology & Biotechnology, Tel-Aviv University, Ramat Aviv 69978, Tel-Aviv, Israel
The availability of monoclonal antibodies (mAbs) which bind to a specific antigen at distinct and well-defined sites led to a better understanding of the effect of highly specific antigen-antibody interactions on antigen behavior. We found that appropriate mAbs interact with their antigens at sites where unfolding is started, leading to stabilization, refolding and suppression of further aggregation. These effects were found to be related to the localization of the antigen site of the antibody. In the following we study the interaction of carboxypeptidase A (CPA) and Alzheimer's 13-amyloid peptide (~AP) with some of their selected monoclonal antibodies. From a large panel of mAbs we selected two mAbs which exhibit significant chaperone-like activity in the refolding and prevention of CPA aggregation induced by heat and excess of Zn ions. CPA exposed to aggregation conditions maintains its solubility and catalytic properties in the presence of mAbs CP10 and CP 9. In vitro aggregation of ~AP (1-40) was inhibited by the presence of one of its mAbs, called AMY-33, but not by mAb 6F/3D, reinforcing the dependence of prevention of aggregation on the localization of aggregating epitopes. Identification of sequences or regions that may play a role in the protein folding pathway via appropriate mAbs may lead to understanding and prevention of protein aggregation.
1. INTRODUCTION
Protein aggregation is a major problem that extends into the mechanisms of human diseases and the fundamental aspects of protein folding, expression and function. Data from several laboratories(1-3) suggest that aggregation is non-specific in the sense that addition of other proteins can influencethe extent of aggregation of a certainprotein. The availabilityof monoclonal antibodies has facilitatedunderstanding how highly specific antigen-antibody interactions affect antigen stability. The complementary conformation between the interactingregions of the antibody and its antigen may confer high specificityand stabilityto the immunocomplex formed (4). Like the ubiquitous chaperones (5-7) mAbs raised against specific,native antigens may assistin antigen refolding (8-10) by recognizing incompletely folded epitopes and inducing their native conformation. By appropriate selection,mAbs have been found to bind to predefined locations on protein or peptide antigens without inhibitingtheirbiological activity(11). The identificationof such classes of sequences that participatein folding-unfolding and/or solubilization-aggregation
184 processes suggests the existence of effective solutions for the prevention of aggregation (1214). Interesting results were obtained from the study of the interaction of carboxypeptidase A (CPA) and/or Alzheimer's 13-amyloid peptide (13AP) with some of their selected monoelonal antibodies. CPA, a well-characterized zinc-exopeptidase that exhibits both peptidase and esterase activity, occupies a prominent position in the bibliography of metalloenzymes (15). A large number of mAbs have been prepared towards native enzyme and their properties have been widely investigated (11,16). Some of these antibodies bind to the enzyme with a relatively high binding constant at a location remote from the enzyme's active site and assist in refolding of heat-denatured enzymes (10). From a large panel of mAbs we selected two mAbs which which were found to exhibit significant chaperone-like activity in the prevention of CPA aggregation induced by heat and excess of Zn ions (17). In the other antibody-antigen complex we found that in vitro aggregation of 13AP (140) was inhibited by the presence of one of its mAbs, called AMY-33, but not by mAb 6F/3D, reinforcing the dependence of prevention of aggregation on the localization of aggregating epitopes. ELISA measurements and determination of residual enzymic activity as a probe of monitoring the native structure are used in our study to follow the inhibitory effect of different mAbs on prevention of aggregation of CPA and/or 13AP.
2. MATERIALS AND METHODS
2.1. Carboxypeptidase A aggregation CPA was obtained as an aqueous crystalline suspension (Sigma Chemical Co.). The crystals were washed with double-distilled water, centrifuged, and dissolved in 0.05 M Tris/HC1/O.5 M NaC1 buffer, pH 7.5. Protein concentration was determined by UV absorbance at 278 nm and by the Bradford method using BSA as a standard (18). The enzymic activities of CPA and its immunocomplexes were determined spectrophotometrically at 254 nm, using hippuryl-DL-13-phenyl-lactic acid as esterase substrate in 0.5 M PBS/0.5 M NaCl buffer, pH 7.5 (19). The preparation and characterization of the mAbs CPl0 and CP 9 chosen for the present study were described previously (16, 20). The optimal denaturation temperature of 50~ was chosen for CPA, because at this temperature the enzyme is easily denatured and the relevant mAbs have been shown to retain their binding affinity (21). The effect of immunocomplexation between CPA and its mAbs on enzyme aggregation was monitored by determination of the residual enzymic activity of CPA, as follows. CPA (0.1-1 ~tg) in PBS in the presence or absence of excess of ZnC12 was measured after exposure to aggregation conditions by incubation at 50~ with or without mAbs CPt0 and CP 9 (100 ~tl in PBS), at antibody/CPA molar ratios l:l. The enzymic activities of the immunocomplexes formed were measured as described above. Data are expressed as the percentage of activity reported to the enzymic activity of CPA before denaturation (being considered 100%).
185
2.2. 13-Amyloid Peptide aggregation Synthetic [3AP (1-40) was obtained from Sigma. For in vitro induced aggregation, the reaction mixture tubes containing 200 ~tl of an aqueous solution of [~AP (2.5 x 10-5 mM) were incubated for 3 h at 37~ Aggregated [3-amyloid samples were removed by centrifugation for 15 min at 15,000 x g. To determine the residual soluble ~AP, the supernatants were then incubated for another 60 min with an excess of mAb AMY-33 and/or 6F/3D to produce immunocomplexed [3AP. The two commercial mAbs used in this study, raised against peptides 8-17 and 1-28 respectively of [3AP, were anti-human [3-amyloid mAb 6F/3D (Accurate Chemicals, Westbury, NJ, USA) and mAb AMY-33 (Zymed, San Francisco, CA, USA). Rabbit polyclonal antibodies raised against synthetic ~AP-(1-40) were obtained from Boehringer Mannheim, Germany. In another set of experiments mAbs, at equimolar antibody/antigen concentrations, were added to the reaction mixtures before the first incubation period of 3h at 37~ The amount of 13AP left in solution under the various conditions was measured by ELISA as follows. Rabbit anti-[3-amyloid (1-40) antibody (100 ng/well) was attached to epoxy-coated Microtiter plates for 16 h at 4~ Atter the plates were washed with phosphate-buffered saline (PBS) containing 0.005% Tween 20, the residual epoxy groups were blocked by adding 1% low-fat milk. Before use, the plate was thrice washed with PBS/Tween 20 and then dried. The soluble immunocomplex of anti-13-amyloid/13AP obtained as described above was added to the plates for an additional 1 h at 37~ and bound mAb was measured by horseradish peroxidase-labeled goat-anti-mouse antibody. Degradation of the O-phenylenediamine substrate by HRP was monitored at A495 according to manufacturer's instructions by using an ELISA reader. The amount of mAb bound was assumed to be proportional to the amount of soluble amyloid peptide that remained in the reaction tube after incubation at the various aggregation conditions specified. The data represent the mean of three replicates. The standard deviations of the intra-assay and inter-assays were <5% in all cases. 3. RESULTS
3.1. Effect of immunocomplexation on the in vitro aggregation of Carboxypeptidase A The immunoeomplex formation led to prevention of aggregation of CPA induced by heat or metal ions. The immunoeomplexation of CPA with some of its selected mAbs maintained not only the solubility of the protein but also its enzymic activity. Even a great excess of unrelated monoclonal antibodies did not affect the aggregation of CPA molecules induced under the same experimental conditions (Fig. 1A,B).
186
Figure 1. Protective effect of monoclonal antibodies on the prevention of CPA aggregation. A) The enzyme was exposed to 50~ C for 1 h and residual enzymic activity was measured as described in Experimental section. The enzyme activity of the following reaction mixtures (1) CPA alone, (2) CPA + mAb CP9, (3) CPA + mAb CPl0, (4) CPA + unrelated antibody were compared and related to enzymic activity of the same amount of enzyme before denaturation. B) The experiment was repeated in the presence of excess of ZnCI2 with the same reaction mixtures (2) to (4).
3.2 Effect of immunocomplexation on the in vitro aggregation of [3-Amyloid Peptide. Monoclonal antibodies were added to the reaction mixture before or after exposure of synthetic 13-amyloid peptide to aggregation conditions in the presence of heparan sulfate and/or metal ions Zn 2+ and A13+ (22). The results show (Fig. 2A) that mAb AMY-33, which recognizes an epitope spanning amino acid residues 1-28 of 13AP, inhibited the aggregation of peptide in the presence or absence of heparan sulfate. No inhibitory effect on metal-induced amyloid aggregation was seen under the same experimental conditions. The mAb 6F/3D, which recognizes an epitope located between residues 8 and 17 of IAP, slightly interfered with Zn 2+ -induced aggregation but had no effect on the self-aggregation induced by other aggregation-inducing agents (Fig. 2B), suggesting different aggregation epitopes involved in each reaction.
187
Figure 2. Effect of immunocomplexation on the in vitro aggregation of J3AP. Relative amounts of residual soluble 13AP (1-40) in the absence (-) or in the presence (+) of mAbs AMY-33 (A) or 6F/3D (B), as measured by ELISA (as described in Materials and Methods). 1.J3AP; 2. [3AP plus 50 mM heparan sulfate; 3. J3AP plus 103M A1C13; 4. ~AP plus 103M ZnC12. 4. DISCUSSION
Antibodies may be used as reporting probes for the detection of antigens and conformational changes induced by various external factors. Moreover, they play an active role in inducing changes in the antigen molecule. Highly specific mAbs may act by changing the molecular dynamics of the antigen molecule and may induce structural rearrangements in the molecular edifice.
188 The data presented show that the antibodies exhibited an inhibitory effect on the antigen aggregation which was related to localization of the antibody binding sites and to the nature of the aggregating agents. Monoclonal antibodies are able to recognize incompletely folded epitopes and induce native conformation (8-10), to stabilize the antigen against denaturing factors with prevention of aggregation (21,22) and to resolubilize already formed protein aggregates (23). The active role of mAbs requires a high binding constant to the strategic positions on the antigen molecule and to be non-inhibitory to the biological activity of the respective antigen. The identification of the "aggregating epitopes" as sequences related to the sites where protein aggregation is initiated and preparation of mAbs against these regions may lead to understanding and prevention of protein aggregation. The data available in literature suggest that for practically all the antigens it might be possible to prepare monoclonal antibodies which bind to preselected epitopes with a high affinity to the antigen without affecting their catalytic activity. At least 15 different polypeptides are known to be capable of causing in vivo different forms of pathological amyloidosis via their deposition in particular organs or tissues as insoluble protein fibrils. The formation of immunocomplexes with such highly specific monoclonal antibodies may provide a general and convenient method to prevent in vivo aggregation of such proteins.. REFERENCES
1. L.R. De Young, A.L. Fink and K.A. Dill, Acc. Chem. Res. 26 (1993) 614. 2. R. Wetzel, Trends Biochem. Sci. 12 (1994) 193. 3. R. Wetzel, J. Perry and C.Veilleux, Bio/Technol. 9 (1991) 731. 4. M.E. Goldberg, Trends Biochem. Sci. 16 (1991) 358. 5. R.J. Ellis and S.M.Van der Vies, Ann. Rev. Biochem. 60 (1991) 321. 6. M.J. Gething and I. Sambrook, Nature 355 (1992) 33. 7. J.P. Hendrick and F.U. Hartl, Ann. Rev. Biochem. 62 (1993) 349. 8. S. Blond and M. Goldberg, Proc. Natl. Acad. Sci. USA 84 (1987) 1147. 9. J.D. Carlson and M.L. Yarmush, Bio/Technology. 10 (1992) 86. 10. B. Solomon and F. Schwartz, J. Molec. Reeog. 8 (1995) 72. 11. B. Solomon, N. Moav, G. Pines and E. Katchalsky-Katzir, Molec.Immunol. 21 (1984) 1. 12. J.L. Silen and D.A. Agard, Nature 341 (1989). 462. 13. X. Zhu, Y. Ohta, F. Jordan and M. Inouye, Nature 339 (1989) 483. 14. J.R. Winter, P. Sorensen and M.C. Kielland-Brandt, J. Biol. Chem. 269 (1994) 22007. 15. B.L. Vallee and A. Galdes, Adv. Enzymol RAMB. 56 (1984) 283. 16. B. Solomon, R. Koppel, D. Kermet and G. Fleminger, Biochemistry 28 (1989) 1235. 17. T. Katzav-Gozansky. M.Sc. Thesis. Tel-Aviv University (1995). 18. M. M. Bradford, Anal. Biochem. 72 (1976) 248. 19. J.R. Whitaker, R. Menger and M.L. Bender, Biochemistry 5 (1966) 386. 20. B. Solomon and N. Balass, Biotech. Appl. Biochem. 14 (1991) 202. 21. T. Katzav-Gozansky, E. Hanan and B. Solomon, Biotech. Appl. Biochem 23 (1996) 227. 22. B. Solomon, R. Koppel, E. Hanan and T. Katzav-Gozansky. Proc. Natl. Acad. Sci. USA 93 (1996) 452. 23. B. Solomon, R. Koppel, D. Frenkel and E. Hanan-Aharon, Proc. Natl. Acad. Sci. USA 94 (1997) 4109.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
189
Stability of m o n o d e a m i d a t e d forms of r i b o n u c l e a s e A F. Catanzano a, G. Graziano b, S. Capasso c and G. Barone a aDept. Chemistry, University "Federico II", Via Mezzocannone, 4 - 80134 Naples, Italy bFaculty of Science, University of Sannio, Via Marmorale - 82020 Paduli (BN), Italy CFaculty of Environmental Science, Second University of Naples Via Arena, 22 - 81100 Caserta, Italy
The effect of selective deamidation of bovine pancreatic ribonuclease A, RNase A, on its thermal stability has been investigated by means of differential scanning calorimetry at changing the pH. Two isomers are obtained in which asparagine 67 is replaced by an aspartyl residue or an isoaspartyl residue. The pH values are limited to the range in which the unfolding process is reversible. The monodeamidated forms have a different thermal stability with respect to the parent enzyme. In particular, the replacement of asparagine 67 with the isoaspartyl residue, involving the insertion of a methylene group in the external loop Cys65Cys72, leads to a substantial decrease in thermodynamic stability.
1. INTRODUCTION Deamidation of asparagine and glutamine side-chains frequently occurs during purification of natural and recombinant proteins [1]. This is of biotechnological interest concerning the preparation, storage and trading of such products. The reaction is also observed in vivo as a post-translational modification, and is most likely related to protein aging [2]. The reaction mechanism depends on pH: i) at acidic conditions the direct hydrolysis of amidic side-chains is dominant, leading to the formation of a normal aspartyl residue, n-Asp; ii) at neutral and basic conditions the reaction occurs via the so-called "[3-shift" mechanism, consisting of a nucleophilic attack on the carbonilic carbon of the asparagine side-chain by the peptide nitrogen of the subsequent amino acid residue in the chain [1,3]. The intramolecular reaction proceeds via a cyclic succinimidyl intermediate whose spontaneous hydrolysis can occur on both sides of imide nitrogen giving rise to two products: one containing an aspartyl a-linked, n-Asp, and the other containing an aspartyl 13-1inked to the adjacent residue in the main chain, iso-Asp (see Figure 1). In this case a methylene group is transferred from the sidechain into the backbone, and the reaction rate is strongly influenced by primary structure: the higher deamidation rates are shown by asparagine residues followed by glycine, both in
190 peptides and proteins [1,3]. On the other hand, the three-dimensional structure also plays a fundamental role, as it imposes conformational constraints on the sites accessible to deamidation [4]. 0 II
/ ~He ENI4H|
./rH
--- NN
/ t
O
0
II
12-.Oi4
~'~'E--NH- EH--EO-II I 0
Asn-peptide
E~NH--EH~EO-II
0
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,~r'H -NIl
~
"- - NU/
~-Asp-peptide
II
I N-rH--m--~ ~"' EH-~,,r/ I /E. n
0
-HN
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E-- Oil II
0
13-Asp-peptide
Figure 1. Deamidation of asparagine via the "[3-shift" mechanism. The effect of deamidation on the thermal stability of RNase A, in a wide range of pH, has been studied by means of DSC measurements. This enzyme can be selectively deamidated in vitro at a single residue, namely asparagine 67, under mild conditions. In fact, it is possible to isolate a chemically pure form of two monodeamidated derivatives: (N67D)RNase A containing n-Asp at position 67, and (N67isoD)RNase A containing iso-Asp at position 67 [5]. Furthermore, the three-dimensional structure of (N67isoD)RNase A has been recently determined to 1.9 A by X-ray crystallography [6]. This makes it possible to establish a correlation between thermodynamic findings and structural features. It is found that (N67isoD)RNase A is markedly less stable than the parent enzyme: the denaturation temperature and enthalpy change actually decrease by 6.3 ~ and 65 kJ mo1-1, respectively, with respect to RNase A at pH 5.0. The structural comparison between the two proteins leads to the conclusion that the difference in thermal stability is due to the loss of two hydrogen bonds in the loop containing residue 67 of (N67isoD)RNase A.
2. MATERIALS AND METHODS Selective deamidation of RNase A is carried out following the procedure devised by D'Alessio and co-workers [5]. Two monodeamidated forms are obtained: one containing an aspartyl residue at position 67, and the other containing an isoaspartyl residue at position 67, both of which had a more negative net charge than the unreacted RNase A. The mixture of the two monodeamidated proteins is separated from unreacted RNase A and polydeamidated forms by means of ionic exchange chromatography, using a carboxymethyl cellulose resin.
191 The monodeamidated fraction of RNase A is collected, dialyzed and lyophilized. The two monodeamidated proteins are separated by means of HPLC with a reverse-phase chromatography, taking advantage of their different hydrophobicity. The ratio of the two eluted peaks is approximately equal to 1.5/1.0, and the more abundant fraction corresponds to the protein containing the isoaspartyl residue. Conservative cuts are made to ensure preparations of high purity. The two monodeamidated proteins, (N67isoD)RNase A and (N67D)RNase A, are exhaustively dialyzed to completely remove (NH4)2SO4, lyophilized and stored at-18 ~ for successive investigations. The protein concentration of dialyzed samples is determined spectrophotometrically, using the extinction coefficient of RNase A at 278 nm, 9800 M -j cm -1 for all proteins, because the cromophore groups are not affected by deamidation. Buffers are: at pH 3.0 glycine-HCl; at pH 4.0 and 5.0 acetic acid-sodium acetate; at pH 6.0 MES. DSC measurements are performed and analyzed as described elsewhere [7]. Before DSC scans samples are dialyzed against the required buffer at 4 ~ for 24 hours.
3. RESULTS Thermal stability of RNase A and the two monodeamidated proteins is characterized in the pH range 3.0-6.0 and the results are summarized in Table 1. Although the protein concentration is not a significant variable of the process, all DSC measurements are performed at [P] = 0.140 mM and the buffer concentration is 0.1 M. At all investigated pHs the process proves reversible, according to the reheating criterion, and the value of the cooperative unit, CU = AdH(Td)/AdHv.H.(Td), is always close to one. Therefore, the temperature-induced denaturation of the three proteins is well represented by the two-state N r D transition model, under all conditions investigated. The values of denaturation temperature decrease significantly on lowering pH from 5.0 to 3.0: Td passes from 61.3 to 52.4 ~ for RNase A, from 61.5 to 49.0 ~ for (N67D)RNase A, and from 55.0 to 43.2 ~ for (N67isoD)RNase A. The decrease is more significant for the two monodeamidated forms with respect to the parent enzyme. Indeed (N67D)RNase A, whose denaturation temperature is very close to that of RNase A at pH 5.0, shows T d values smaller than RNase A on lowering pH (i.e., 55.3 ~ against 57.0 ~ at pH 4.0, and 49.0 ~ against 52.4 ~ at pH 3.0). Furthermore, the difference in Td between RNase A and (N67isoD)RNase A rises from 6.3 ~ at pH 5.0, to 7.5 ~ at pH 4.0 and to 9.2 ~ at pH 3.0. On the other hand, an increase of pH from 5.0 to 6.0 causes only a minor rise in denaturation temperature for all proteins. Finally, the increase of denaturation enthalpy on raising pH from 3.0 to 6.0 for all proteins can be directly ascribed to the increase of T d and to the large positive value of AdCp. The latter parameter is practically not affected by the selective deamidation of Asn67, as expected on the basis of models developed for calculating AdCp [8], and amounts to about 5.5 kJ K-lmo1-1. The strong difference between the two monodeamidated proteins is quite unexpected. However, they differ for the insertion of a methylene group at position 67 in the backbone of (N67isoD)RNase A, and this structural modification can strongly affect thermal stability.
192 Table 1. Thermodynamic parameters of the thermal denaturation process of the three ribonucleases at different pH values from DSC scans. Protein concentration is 0.140 mM (1.9 mg mL "1) and buffer concentration is 0.1 M in all the measurements.
RNase A
(N67D)RNase A
(N67isoD)RNase A
pH
Td (~
AdH(Td) (kJ mol -!)
AdS(Td) (kJ K-lmo1-1)
AdC p (kJ K-lmo1-1)
CU
3.0
52.4
415 • 13
1.27 • 0.04
5.0 • 0.6
0.99
4.0
57.0
440 • 12
1.33 • 0.04
5.4 • 0.5
0.99
5.0
61.3
465 • 13
1.39 • 0.04
5.5 • 0.6
1.01
6.0
62.4
470 • 14
1.40 • 0.05
5.8 • 0.6
1.02
3.0
49.0
390 + 12
1.21 + 0.04
5.8 + 0.7
0.99
4.0
55.3
430 • 14
1.31 • 0.05
5.4 + 0.6
1.01
5.0
61.5
460 • 14
1.37 • 0.04
5.5 • 0.6
1.03
6.0
62.2
465 • 13
1.39 • 0.04
5.2 • 0.5
1.01
3.0
43.2
335 • 10
1.06 • 0.04
5.6 • 0.6
1.01
4.0
49.5
370 • 12
1.15 • 0.04
5.1 • 0.5
0.98
5.0
55.0
400 • 10
1.22 • 0.03
5.4 • 0.6
0.98
6.0
55.8
405 • 11
1.23 • 0.03
5.8 • 0.6
1.02
Note. Each figure is the mean value of at least four measurements. The error in T d does not exceed 0.2 ~ Reported errors for AdH(Td) and AdCp are the standard deviations of the mean from the multiple determinations. Reported errors for AdS(Td) are calculated by propagating the errors for AdH(Td) and T d.
According to electrostatic principles, the maximum stability of globular proteins should occur at their isoelectric point. The isoelectric point of RNase A is about 9.3, while for both monodeamidated derivatives it is lower due to the additional negative charge. On this basis, it is conceivable that there is a different pH dependence of the thermal stability for RNase A and the two deamidated forms. By plotting the AdH(Td) values versus T d values it results that there is no difference in enthalpy between (N67D)RNase A and the parent protein, whereas there is a marked difference with respect to the derivative containing the isoaspartyl residue [7]. Therefore, the decrease in thermal stability on lowering pH from 5.0 to 3.0 of (N67D)RNase A with respect to RNase A must be due to entropic effects, in line with general results obtained by means of theoretical calculations [9].
193
4. DISCUSSION The significantly lower thermal stability of the protein containing the isoaspartyl residue with respect to RNase A merits a deepening. The thermodynamic analysis is restricted to measurements at pH 5.0, even though very similar conclusions are reached by selecting any other pH in the range 3.0-6.0. The stability curves, AdG vs T, which are a measure of the work required to destroy the native conformation [ 10], can be calculated according to the following equation: (1)
AdG(T ) = AdH(Td)[1 - (T/Td) ] + AdCp[T - T d - T ln(T/Td) ]
which is exact in the assumption that AdC p is temperature-independent. The stability cu rv es , with the characteristic parabolic-like profile, of the two proteins at pH 5.0 are reported in Figure 2" the temperature corresponding to the maximum is not strongly affected by the N67isoD substitution, occurring for both proteins around 260 K. In contrast, the denaturation Gibbs energy change of (N67isoD)RNase A is significantly lower than that of the parent enzyme at any temperature. It is important to evaluate the enthalpy and entropy contributions determining such a decrease in thermodynamic stability.
0
,
,
,
,
,
,,,
,
,
200
220
240
260
280
300
320
5O Q
,,.-,
E
40
3O r~ 20
10
o lao
!
340
temperature ( K ) Figure 2. Stability curves of (N67isoD)RNase A (curve a) and RNase A (curve b) at pH 5.0, calculated using the parameter values reported in Table 1. For a correct comparison, AdH and AdS have to be known at the same temperature. The T d value of (N67isoD)RNase A, 55.0 ~ is selected as reference, to minimize
194 extrapolation errors. For a two-state transition, assuming AdCp temperature-independent, the denaturation enthalpy and entropy can be calculated according to the well-known equations: AdH(T) = AdH(Td) + AdCp 9(T- Td)
(2)
AdS(T ) = [AdH(Td)/Td] + AdCp 9In(T/Td)
(3)
Performing the calculations for RNase A, it results AdH(55 ~ = 430 • 13 kJ mo1-1 and ADS(55 ~ = 1.29 + 0.04 kJ K-lmo1-1. These figures are 30 kJ mo1-1 and 70 J K-lmo1-1, respectively, greater than the corresponding ones of the modified protein. The Gibbs energy difference between RNase A and (N67isoD)RNase A amounts approximately to 7.0 kJ mol -l at 55 ~ As a decrease in AdH has a destabilizing effect, whereas a decrease in AdS stabilizes the folded structure, the analysis leads to the conclusion that the destabilization of (N67isoD)RNase A with respect to the parent enzyme is due to enthalpic factors, partially compensated by entropic factors. It is possible to correlate the thermodynamic results to the structural data. Asparagine 67 in RNase A is located in a type I 13-tum of an external loop constituted by 8 residues and closed by the disulfide bond between Cys65 and Cys72 [11]. Three hydrogen bonds are important in determining the conformation of the loop: i) that between O of Cys65 and N of Gly68 with a bond length of 2.80 A; ii) that between O 8 of Asn67 and N of Gin69, which gives rise to a structure resembling a C l0 [5-turn; iii) that between O of Gln69 and N of Cys65 with a bond length of 2.62/1,. It is worth noting that these hydrogen bonds were found not only in the X-ray structure of RNase A, but also in the NMR solution structure of the enzyme [ 12]. According to the X-ray diffraction results by Mazzarella and co-workers [6], the threedimensional structure of (N67isoD)RNaseA is very similar to that of the parent enzyme, except that the loop Cys65-Cys72 undergoes a substantial conformational change, with a shift toward the main body of the molecule. The insertion of an additional methylene group in the main chain, with formation of a 13-carboxylic bond, causes a significant enlargement of the loop and the loss of the hydrogen bond characteristic of the ~l-turn: the distance between O of Cys65 and N of Gly68 is now equal to 4.40 A. Obviously, also the hydrogen bond involving the side-chain oxygen of Ash67 is lost. On the other hand, the hydrogen bond between O of Gin69 and N of Cys65 is conserved, even though the bond length is increased to about 3.0 A. Moreover, the loop Cys65-Cys72 is in contact with the active site in RNase A, by means of two hydrogen bonds involving the side-chain of Asp l21 [ 11 ]: i) that between 0 81 of Aspl21 and N of Asn67, which is mediated by a water molecule; ii) that between 0 82 of Aspl21 and N of Lys66. In the structure of (N67isoD)RNase A, Aspl21 does not change its conformation and the two hydrogen bonds are conserved [6]. In particular, as a consequence of the shift toward the main body of the molecule experienced by the loop, the hydrogen bond between 0 81 of Asp l21 and N of isoAsp67 is direct. Therefore, it can be concluded that the N67isoD substitution causes the loss of two important hydrogen bonds in the external loop
closed by the disulfide bridge between Cys65 and Cys72, whereas the overall structure of the enzyme is not altered. The loss of two hydrogen bonds correlates fairly well to the observed decrease in thermodynamic stability. Indeed, as strongly advocated by Pace and co-workers [13], intramolecular hydrogen bonds greatly contribute to the stability of native conformation. The comparison between RNase A and (N67isoD)RNase A shows that each hydrogen bond contributes about 3.5 kJ mo1-1 to the Gibbs energy of protein stabilization at 55 ~ which is in agreement with estimates from mutational studies [ 13]. Additionally, the data point out that hydrogen bonds are enthalpically stabilizing. On the other hand, the increased flexibility of the extemal loop, due to the insertion of an extra methylene group in the main chain and the absence of two hydrogen bonds, agrees with the observed decrease in denaturation entropy of (N67isoD)RNase A with respect to the parent enzyme. In fact, locally, the folded structure of the modified protein is less conformationally rigid. Thus, the partial compensation between the destabilizing enthalpic contribution and the stabilizing entropic one proves clear: any loosening of intramolecular interactions in the folded conformation tends to be compensated by a gain of degrees of freedom. A detailed comparison of the water sites close to the external loop between the crystal structure of (N67isoD)RNase A and that of RNase A has not been carried out hitherto. Changes in water of hydration could have effects on the protein stability. However, Palmer and co-workers [ 14], found that only 17 water sites, out of 100-200 water molecules identified in X-ray structures, are conserved in RNase A structures from 5 different space groups. None of these 17 conserved water molecules is bound to residues in the loop Cys65-Cys72. This suggests that, being the loop well exposed to the solvent, the water molecules of hydration are not closely associated with protein atoms. According to the analysis by Dunitz [15], the entropy cost of transfcrring a water molecule from the liquid to the protein falls in the range of between 0 and 30 J K-lmo1-1, at room temperature. The upper limit will apply only to those few water molecules that are most firmly bound (i.e., those having Debye-Waller temperature factors of around 5 to 10 A2). Most of the remaining water content of a protein, including much of the surface water, is likely to cost only a small fraction of this upper limit and will behave more like liquid water. This discussion leads to the conclusion that the role played by changes in water of hydration in causing the lower thermal stability of (N67isoD)RNase A with respect to the parent en~me should be negligible. The loss of two intramolecular hydrogen bonds proves to be a convincing rationale for the destabilization caused by the N67isoD substitution. In fact, the insertion of an additional methylene group into the polypeptide backbone is a more drastic modification than other amino acid substitutions affecting only side-chains. In addition, it occurs in a region of the protein sequence considered as a possible chain-folding initiation site: the first preferred disulfide pairing should occur between Cys65 and CysT2, because hydrogen bonds among neighbour residues play a fundamental role for the formation of locally ordered structures along the overall unfolded polypeptide chain, as stressed by Scheraga and co-workers [ 16]. On this basis, it is conceivable that (N67isoD)RNase A shows different physico-chemical and
196 biological properties. Indeed, (N67isoD)RNase A, after denaturation and reduction of disulfide bridges, refolds at a significantly lower rate than RNase A and (N67D)RNase A [5]. Moreover, it possesses different Michaelis-Menten constants for several substrates compared with those of RNase A and (N67D)RNase A [5], and a binding affinity for 2'CMP which is greater than that of the other two proteins [7]. In conclusion, the selective deamidation of Asn67 in RNase A, giving rise to two monodeamidated derivatives, containing an aspartyl residue or an isoaspartyl residue at position 67, causes a decrease in the protein stability. In particular, for (N67isoD)RNase A, the destabilization agrees fairly well with the structural consequences of the methylene insertion in the main chain of the external loop Cys65-Cys72. These thermodynamic results seem to support the idea that deamidation plays a physiological role in controlling the in vivo half-life of proteins, by lowering their thermodynamic stability. Acknowledgments. Work supported by P.R.I.N. grant from the Italian Ministry for University and Scientific and Technological Research (M.U.R.S.T., Rome) and University of Naples "Federico Ir'.
REFERENCES
1. H.T. Wright, Crit.Rev.Biochem.Mol.Biol., 26 (1991) 1. 2. A. Di Donato, P. Galletti and G. D'Alessio, Biochemistry, 25 (1986) 8361. 3. H.T. Wright, Protein Eng., 4 (1991) 283. 4. S. Clarke, Int.J.Pept.Protein Res., 30 (1987) 808. 5. A. Di Donato, M.A. Ciardiello, M. de Nigris, R. Piccoli, L. Mazzarella and G. D'Alessio, J.Biol.Chem., 268 (1993) 4745. 6. S. Capasso, A. Di Donato, L. Esposito, F. Sica, G. Sorrentino, L. Vitagliano, A. Zagari and L. Mazzarella, J.Mol.Biol., 257 (1996) 492. 7. F. Catanzano, G. Graziano, S. Capasso and G. Barone, Protein Sci., 6 (1997) 1682. 8. G. Graziano, F. Catanzano, P. Del Vecchio, C. Giancola and G. Barone, Gazz.Chim.It., 126 (1996) 559. 9. D. Stigter and K.A. Dill, Biochemistry, 29 (1990) 1262. 10. W.J. Becktel and J.A. Schellman, Biopolymers, 26 (1987) 1862. 11. A. Wlodawer, L.A. Svensson, L. Sjolin and G.L. Gilliland, Biochemistry, 27 (1988) 2705. 12. J. Santoro, C. Gonzales, M. Bruix, J.L. Neira, J.L. Nieto, J. Herranz and M. Rico, J.Mol.Biol., 229 (1993) 722. 13. J.K. Myers and C.N. Pace, Biophys.J., 71 (1996) 2033. 14. I. Zegers, D. Maes, M. Dao-thi, F. Poortmans, R. Palmer and L. Wyns, Protein Sci., 3 (1994) 2322. 15. J.D. Dunitz, Science, 264 (1994) 670. 16. G.T. Montelione and H.A. Scheraga, Acc.Chem.Res., 22 (1989) 70.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998Elsevier Science B.V. All rights reserved.
197
P r e s s u r e effects on p r o t e i n oligomeric d i s s o c i a t i o n C. Balny INSERM, Unit~ 128, 1919, route de Mende, 34293 Montpellier, Cedex 5, France Abstract High hydrostatic pressure induces changes in protein c o n f o r m a t i o n , solvation and enzyme activities via reversible or non-reversible effects on intraand inter-molecular interactions (noncovalent bonds) [1]. Generally, p r e s s u r e acts by destabilizing hydrophobic bonds which is associated with negative volume changes, inducing dissociation of oligomeric structures. If this hypothesis is accepted, opposite examples exist and some of which are described below. Results are also discussed in terms of possible biotechnological applications where the pressure p a r a m e t e r , associated or not with the temperature p a r a m e t e r , is an i m p o r t a n t factor to achieve protein d e n a t u r a t i o n [2-4].
1. I N T R O D U C T I O N Elevated pressure(*) is now becoming an useful tool to study both structural and functional aspects of proteins. This p a r a m e t e r is unique in its ability to provide a direct m e a s u r e m e n t of the energy and changes in volume of the dissociation/association processes [1,5]. The behavior of proteins u n d e r high pressure is governed by Le Chatelier's principle : application of p r e s s u r e shifts an equilibrium towards the state t h a t occupies a smaller volume. Closely associated is the principle of microscopic ordering which states that, at constant t e m p e r a t u r e , an increase in pressure increases the degree or ordering of the molecules of a substance. Pressure and t e m p e r a t u r e m i g h t have antagonistic effects but both can cause protein dissociation (or denaturation). The native conformation of a protein exists only over a r a t h e r limited range of both p a r a m e t e r s as shown in Figure 1. Moreover, the effect of pressure on proteins depends on the relative contribution of the i n t r a m o l e c u l a r forces which determine their stability and function. Noncovalent m o l e c u l a r interactions are affected by pressure. Solvation of charged groups is accompanied by volume reduction. Formation of coulombic and hydrophobic interactions are accompanied by positive changes in volume. C h a r g e - t r a n s f e r interactions and stacking of aromatic rings show small decrease in volume and hydrogen bonds are almost pressure-insensitive. The direct consequences are that hydrophobic interactions are destabilized by pressure. * 1 atm = 1 kg.cm 2 = 1.01325 bar = 0.101325 MPa
198 Through these effects, pressure affects protein structure, at the secondary, tertiary and quaternary levels, resulting in protein denaturation. The extent and the reversibility (or irreversibility) of pressure effects depend on the pressure range, the rate of compression, and the duration of exposure to elevated pressures. These effects are also influenced by other e n v i r o n m e n t a l parameters, such as the temperature, the medium composition (pH, salts, solvent). Denatured, AGd
, s"'~
a
is SS
~G~=O
II
~
"~
2-
Native AGd>O ffl m
I~.
1-
.
-20
0
.
.
.
.
.
20
40
60
Temperature (~
Figure 1" Pressure-temperature transition diagram for protein dissociation] denaturation. Adapted from Ref. 4. 2. SPECIFIC METHODS The characterization of dissociation/association reactions usually depends on some average spectroscopic property sensitive to the change in state of association. It is possible to observe changes in fluorescence polarization of an intrinsic or extrinsic probe, changes of center of spectral mass of the intrinsic fluorescence or optical density and energy transfer between fluorophores attached to the subunits. However, in some cases, when no spectroscopic means are available, other methods such as high-pressure polyacrylamide gel electrophoresis have been developed [6]. This method, introduced by Hawley and Weber is a direct method of investigation of the state of association of oligomers. A recent spectroscopic development for indirect absorption observation of the degree of dissociation are the second and fourth derivative spectroscopies in the near-ultraviolet region of proteins [7,8]. This permits one to e n h a n c e selectively spectral changes in the UV absorption of phenylalanine, tyrosine
199 and tryptophan due to solvent access into the protein on unfolding and the solvent polarity which affect the amplitude, position and shape of the second and fourth derivative spectral bands. This method is a suitable tool to evaluate changes of the dielectric constant in the vicinity of the aromatic amino acids in proteins which undergo pressure induced structural changes that may result in dissociation. This approach can give complementary information's to those obtained using the analysis of the conformational drift and hysteresis, a phenomenon firstly shown by Ruan and Weber for the tetrameric enzyme glyceraldehyde phosphate dehydrogenase [9].
3. HIGH PRESSURE AND DISSOCIATION 3.1. Oligomerie dissociation For most of oligomeric enzymes, the exposure to pressure below 100 - 200 MPa often shifts the equilibrium between oligomers towards dissociation into subunits with a lost of activity. This phenomenon is due to the i n t e r s u b u n i t regions of oligomeric proteins which are rich in electrostatic and hydrophobic contacts which stabilize the quaternary structure and which are weakened at relative moderate pressure. This field is very well documented after the pioneering work of G. Weber and coworkers [10]. Detailed studies have been published on the pressure-induced dissociation/inactivation of dehydrogenase, tryptophan synthase and a small dimeric Arc repressor protein. For this protein, phase-sensitive two-dimensional correlated spectroscopy (COSY) and nuclear Overhauser effect enhancement spectroscopy (NOESY) have been used [11]. The intersubunit ~-sheet in the native dimer is replaced by an intramonomer ~-sheet in the denatured state at a pressure higher t h a n 200 MPa with formation of a denatured molten globule monomer. A recent example on yeast enolase has been studied in our laboratory by M.J. Kornblatt, using a combination of the fourth derivative ultraviolet spectroscopy under pressure and the high pressure stopped-flow kinetics [12]. Enolase has been characterized as a dimer, each dimer consisting of two active sites appearing to be completely independent. The physical independence of the two active sites might lead one to predict that the monomeric form of enolase would also be active. Since hydrostatic pressure is viewed as a gentle and reversible perturbant, earlier study on pressure-induced denaturation of enolase has been continued to determine whether or not the monomers h a d catalytic activity and to compare dissociation produced by hydrostatic p r e s s u r e with that produced by EDTA and KC1. Typical results are shown in Figure 2 indicating that monomers are inactive. Dissociation of this enzyme involves three processes : d i s r u p t i o n of salt bridges, exposure and hydration of buried surfaces and lost of the divalent cation. The results are discussed in detail elsewhere, showing the role of Mg 2§ [13]. Removing the Mg 2§ from enolase, either by adding EDTA or by preparing apoenzyme, displaces the equilibrium towards monomers and decreases the activation volumes involved. Loss of Mg 2§ contributes to the negative activation volume for dissociation (- 250 ml/mol) and this loss occurs, at least partially, in the transition state for dissociation. Dissociation produces major changes in the conformation of the enzyme as observed by CD experiments [13].
200
1
0
0
0
0
0
0
0
0
0
0
0
0
Q 9
0
0
z,. ,I~
0
0.1
0 C U
0.01
o
~
O--
....
100
150
Pressure, MPa
Figure 2 :Effects of pressure on the quaternary structure and activity on yeast enolase. (o) Enzyme activity measured using high-pressure stopped-flow and expressed as percentage of the 0.1 MPa value, (o) pressure dependence of the percentage of quaternary structure calculated from the fourth derivative spectra. Adapted from Ref. 13. 3.2. N o oligomeric dissociation It has been accepted for some time that high hydrostatic pressure is a rather general way to dissociate molecular assemblies. However, the nature of the intersubunit regions is critical and there are a few exceptions to this behaviour. Such is the case for alcohol dehydrogenases from yeast (YADH) or from Thermoanaerobium brockii (TBADH). Both are tetrameric but have different high pressure responses. YADH activity is continuously inhibited by an increase of pressure whereas TBADH affinity for alcoholic subtrates increases if pressure increases. In the case of these two enzymes, unless the oligomers reassociate very quickly, the activity is not correlated to subunit dissociation (monitored by HPLC after pressurization). It has been suggested that enzymes under pressure undergo a molecular rearrangement which can affect the catalytic efficiency [14]. Another interesting example has been found by J. Frank in our laboratory using methanol dehydrogenase from Methylophaga marina (MDH). Raising pressure to 450 MPa results in a spectral transition detected using the fourth derivative method. Concomitant with a red shift, a high pressure decrease of the derivative amplitude is observed indicating that the tryptophans are on a less polar environment at elevated pressure [8] (see Figure 3). After releasing the pressure, both activity and structural modifications are reversible. The question arises 9 what are the origins of the structural modifications ? MDH is a tetramer with two large and two small subunits. Usually, when pressure promotes oligomeric dissociation a blue shift in the UV spectrum is observed due to the hydration of hydrophobic residues upon exposure to the solvent. The present shift observed is a red shift. On the other
201 hand, previous electrophoretic experiments u n d e r high p r e s s u r e showed that the enzyme m i g r a t e d as a single peak even at 200 MPa w h e r e its activity w a s still m e a s u r a b l e in the electrophoretic gel. This indicates t h a t the enzyme is m a i n t a i n e d in its active t e t r a m e r i c s t r u c t u r e in the p r e s s u r e r a n g e explored [15] (Figure 4).
... " ' - . . 0.05
I MPa~
'"'...
9
.d~',,i
~ ~ 0.4
7,
<~ 0.00
o.2 ~ -0.05
-
* Guanidinum.
,
I
285
,
6M
,
1
~.
~
a
I
29o
.,
,
~
.
!
295
. . . .
0.0
wavelength,nm
Figure 3 " P r e s s u r e effect on the fourth derivative s p e c t r u m of MDH. The original zero order spectrum is shown as a broken line, right h a n d scale. F r o m Ref. 8.
-
I
F
1
600
bar
bar
1 0 0 0 . bar
L..'
1500
bar
2000
bar
Figure 4 : P o l y a c r y l a m i n e gel electrophoretic p a t t e r n s of MDH at different p r e s s u r e s . Total polyacrylamide concentration : 6 %. R u n n i n g buffer : 350 mM ~-alanine/acetic acid, pH 4.5. Runs were carried out at 0.3 mA/gel for 90 min. Continuous curves correspond to proteins. Dashed lines are for e n z y m e activity: the active enzyme zones were shown on gels as colorless zones (dashed line wells). Adapted from Ref. 15
202 In contrast to the majority of the studied oligomeric proteins, the results on MDH were interpreted as a pressure-strengthening of the q u a t e r n a r y structure through an increase of the interaction of aromatic amino acids. The stacking of such residues been known to be reinforced by pressure. In the same way, P. Masson and coworkers have used high pressure to study the denaturation of the tetrameric h u m a n butyrylcholinesterase [16]. They found, using electrophoresis at elevated pressure (up to 200 MPa), that pressure alone did not induced either dissociation or unfolding of the native enzyme. On the other hand, Fourier transform infrared spectroscopy u n d e r high pressure showed that secondary structure changes started at 300 MPa and was complete at 600 MPa. Moreover, pressure induced progressive inactivation of the native enzyme correlated with an irreversible d e n a t u r a t i o n of the tetramer and its subsequent aggregation. In the aggregate, the enzyme has a partially unfolded configuration and it has been suggested that p r e s s u r e promoted exposure of patches of hydrophobic residues. The behavior of oligomeric enzymes under pressure can be yet m ore complicated, as in the case of hydroxylamine oxidoreductase from N i t r o s o m o n a s (HAO). HAO is a multimer of an unknow n number of asubunits containing at least 6 c-hemes and one active site (cytochrome P-460). Kinetic studies at high pressure have indicated that this enzyme undergoes a change in structure during catalysis : Arrhenius plots for reduction of c-hemes with substrate at various pressures are biphasic [17]. Further, the values of activation energies change from positive to negative at 80 MPa, indicating that a conformational change occurred at either a critical temperature or p r e s s u r e . The behavior of this multiheme protein in polyacrylamide gel electrophoresis was studied at pressure up to 300 MPa. Between atmospheric pressure and 150 MPa the enzyme did not undergo structural changes detectable in the microgel system, a results well correlated with the activity data, in the s a m e pressure range. At about 200 MPa, the active form of the enzyme was partially dissociated into smaller subunits. Production of the small molecular weight form in this pressure range may represent the first instance of dissociation to the subunit of the enzyme prior to chemical removal of the c-hemes. At h i g h e r pressure (up to 300 MPa), the enzyme was converted to forms which were irreversibly inactive and having a higher apparent molecular m a s s , suggesting aggregation or denaturation. The two consecutive steps may be related as follows : firstly a discrete pressure-induced conformational c h a n g e (localized unfolding and partial dissociation) could result in the u n m a s k i n g of buried groups which are then able to pair with groups at the subunit contact areas. Secondly, participation of pressure-enhanced interactions (hydrogen bonds or stacking of aromatic rings) may involved in polymerization. This study illustrates the potential value of high pressure electrophoresis for studying dissociation of oligomeric proteins [6]. 4. BIOTECHNOLOGICAL APPLICATIONS We have seen that hydrostatic pressure alters the characteristics of proteins, a phenomenon which can be used to modify the properties of biological materials to preserve or improve their qualities. For many years, this was done primarly for food processing, with new improvements developed during the last decade. However, the phenomenon is complicated. For
203 example, it is known that egg white and several other proteins coagulate or form gels at pressure above 300 MPa. The pressure necessary for aggregation as opposed to oligomeric dissociation phenomena) depends on the type of protein and on the experimental conditions (temperature, salts, pH, etc.). It is likely that pressure induces extensive unfolding of protein chains, thus permitting non-native state subsequent intermolecular interaction between exposed aminoacid residues. When these non-covalent interactions are mainly hydrophobic, it is likely that gelation takes place only upon pressure release. I n addition, the role of water may be taken in consideration to explain the sensitivity to pressure of sol-gel transitions in protein and polysaccharides. The potential interest of the food industry for these pressurized gels stimulates a lot of basic research in this field [2,3]. Some applications are now emerging such as pressure-induced gel formation of whey proteins [18]. Another application of high pressure is emerging. It concerns the exploitation of the temperature-pressure diagram of the Figure 1 type in association with the water/ices phase diagram. It is known that at 200 MPa, the water is still liquid at a temperature as low as - 20 ~ It becomes then possible to used this property to study cold denaturation of proteins. The combination of pressure and low temperature has been explored to dissociate oligomeric proteins [19], protein-DNA complexes [20] and virus [21]. The group of Silva, recently reported the cold dissociation of allophycocyanin and liver alcohol dehydrogenase [22] and interpreted data in term of decreasing entropy of the studied systems. ACKNOWI,EDGlVIENTS The author thanks Dr. J. Connelly for critical reading of the m a n u s c r i p t and Drs. J. Frank, A.B. Hooper, J.M. Kornblatt, R. Lange, P. Masson and V.V. Mozhaev for stimulating discussions. R~ERENCF~ 1 V.V. Mozhaev, K. Heremans, J. Frank, P. Masson and C. Balny, Proteins : Structure, Function, and Genetics, 24 (1996) 81. 2 C. Balny, R. Hayashi, K. Heremans and P. Masson (eds) High Pressure and Biotechnology, J. Libbey Eurotext/INSERM, Montrouge, France, vol. 224 1992. 3 R. Hayashi and C. Balny (eds) High Pressure Bioscience and Biotechnology, Progress in Biotechnology Serie, vol. 13, Elsevier, 1996. 4 V.V. Mozhaev, K. Heremans, J. Frank, P. Masson and C. Balny, TIBTech, 12 (1994) 493. 5 M. Grol3 and R. Jaenicke, Eur. J. Biochem., 221 (1994) 617. 6 P. Masson, D.M. Arciero, A.B. Hooper and C. Balny, Electrophoresis, 11 (1990) 128. 7 R. Lange, J. Frank, J.-L. Saldana and C. Balny, Eur. Biophys. J. 24 (1996) 277. 8 R. Lange, N. Bec, V.V. Mozhaev and J. Frank, Eur. Biophys. J. 24 (1996) 284. 9 K. Ruan and G. Weber, Biochemistry 28 (1989) 2144. 10 G. Weber, Biochemistry, 25 (1986) 3626.
204 11 12 13 14 15
X. Peng, J. Jonas, J.L. Silva, Biochemistry 33 (1994) 8323. C. Balny, J-L. Saldana, and N. Dahan, Anal. Biochem., 139 (1984) 178. M.J. Kornblatt, R. Lange and C. Balny, Eur. J. Biochem., 251 (1998) 775. S. Dallet and M-D. Legoy, Biochim. Biophys. Acta, 1294 (1996) 15. I. Heiber-Langer, C. Clery, J. Frank, P. Masson and C. Balny, Eur. Biophys. J. 21 (1992) 241. 16 P. Masson, P. Gouet and C. Clery J. Mol. Biol., 238 (1994) 466. 17 C. Balny and A.B. Hooper, Eur. J. Biochem., 176 (1988) 273. 18 D. Foguel and G. Weber, J. Biol. Chem. 270 (1995) 28759. 19 D. Foguel and J.L. Silva, Ptoc. Natl. Acad. Sci. USA, 91 (1994) 8244. 20 D. Foguel, C.M. Teschke, P.E. Prevelige and J.L. Silva, Biochemistry, 34 (1995) 1120. 21 A. Ferrao-Gozales, C.D. Zandonade, F.P. Mariz, J.L. Silva and D. Foguel in High Pressure Resarch in the Biosciences and Biotechnology (K. Heremans, ed.), Leuven Univ. Press, 1997, p. 363.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
205
Effect of high hydrostatic pressure on e n z y m e stability V. AthOs and D. Combes INSA, Centre de Bioing~nierie Gilbert Durand, UMR 5504, LA INRA, Complexe Scientifique de Rangueil, F-31077 Toulouse cedex 4, France.
The influence of different soluble additives (salts and polyols) on the barostability and thermostability of Kluyveromyces lactis ~-galactosidase has been studied. Polyols (glycerol, erythritol, xylitol and sorbitol) more than salts, were shown to act as very effective agents against pressure deactivation. Moreover, an increased thermostability has been observed under high pressure. Indeed, pressure or temperature can both induce an irreversible inactivation of ~-galactosidase. But moderate pressures (50 MPa - 100 MPa) exerted a protective effect against thermal inactivation. For two other ~-galactosidases from Aspergillus oryzae and Escherichia coli, we have shown that pressure could be used to increase noticeably their thermal stability.
1. I N T R O D U C T I O N The use of high hydrostatic pressure in biotechnology has experienced a constant progression over the past few years. The growing interest in using high pressure to treat biological materials lies in its numerous advantages : as a method of sterilisation in the food industry, allowing the organoleptic properties of the products to be preserved (in comparison with a thermal treatment) [1] ; and as an original thermodynamic variable to study the working mechanisms of some biomolecules [2]. Moreover, some advantages of pressure, in comparison with temperature, are : a quasi-instantaneous transmission (isotropic), an action independent of the geometry and identical in every point (isostatic), and low energy inputs required. The behaviour of proteins under high pressure is the centre of interest of many research groups all over the world. The studies of enzyme stability fall as part of this research field. Pressure may act as a denaturing agent, such as temperature [3], or as a stabilizing agent against thermal deactivation, in the range where pressure alone does not inactivate the enzyme [4-5]. The enhancement of enzyme thermal stability under moderate pressures has been often described for enzymes from thermophilic sources [6].
206 Increasing the stability of enzymes agsinst pressure or temperature denaturation is a subject of great concern for biotechnological applications, because of the various advantages which arise by carrying out enzymatic reactions in non-conventional conditions [7]. The thermostabilisation of enzymes by the use of soluble additives has been already widely studied [8]. In this work, we have chosen to study the influence of salts and polyols on Kluyveromyces lactis ~-galactosidase thermo- and barostability. Furthermore, we have combined temperature and pressure effects to evidenced the protective effect of moderate pressures against thermal inactivation. We have extended our study to ~-galactosidases from Aspergillus oryzae and Escherichia coli. 2. E F F E C T OF BAROSTABILITY
SALTS
AND
POLYOLS
ON
THERMO-
AND
Choosing residual activity of enzyme as a criterion for enzyme stability, the effect of additives on K. lactis ~-galactosidase barostability and thermostability has been investigated. The protective effect (PE) of the additives on the enzyme deactivation by pressure or temperature can be defined as a ratio of enzyme half-life in the presence of additives to the half-life of enzyme without any additive. The half-life (tl/2) is defined as the time needed to reduce the initial activity to 50% of its original value. 1.1. I n f l u e n c e o f salts The thermal stability at 50~ for ~-galactosidase was assayed without any additive and the half-life was about 5 min. The influence of four salts on ~-galactosidase thermal stability at 50~ was studied: NaC1, KC1, NaBr and KBr at different concentrations. Figure 1 shows the protective or denaturing effects induced by salts on the enzyme. 4
"
20 15 3-10
'
~
.~ 9 2~
'
/,
1
...........
0
1
2
3
[salts] ( ~ Figure 1. Protective effect of salts on ~-galactosidase at 50~ as a function of their concentration: NaC1 ( *---4 ), NaBr ( ~ - " ), KBr ( ~ - o ) and KC1 (~--o).
207 The pressure stability of the same enzyme at 250 MPa was assayed without any additive and the half-life was 1.6 + 0.1 rain. Table 1 shows the protective effects induced by salts on the enzyme for a pressure treatment at 2500 bar. Table 1. Protective effect of salts on 6-galactosidase at 2500 bar [salts] (M) KC1 SaC1 KBr 0 li0.1 1__+0.1 1__+0.1 0.5 3__+0.3 0.1__+0.06 l!-0.3 1 5__+0.8 0.1__+0.06 0.3__+0.08 2 14+__2.4 0.2__+0.07 0.3__+0.08 3 60!-6.8 3__+0.5 0.5~0.2 i
l lll
i
NaBr 1__+0.1 0.03!0.01 0.01 0 0
In both cases, NaBr and KBr have a destabilising influence (PE
The influence of the four polyols containing three to six carbon atoms (glycerol (C3), erythritol (C4), xyhtol (C5) and sorbitol (C6)) on ~-galactosidase thermal deactivation has been studied at 50~ and at different polyol concentration. Figure 2 shows the evolution of the protective effect of these additives as a function of their concentration. Erythritol, whatever the concentration, exhibits a destabilising effect. Besides this particular behaviour, the enhancement of the thermal stability caused by the three other polyols is proportional to their molecular size, being glycerol < xylitol < sorbitol. This ranking is verified at any concentration, and the stabilizing effect is increasing with the concentration. It may be seen from the insert of Figure 2 that the protective effect obtained by using a 2M sorbitol solution is near to 300. This is a particularly strong influence, compared with the protective effect of a 2M solution of xylitol (PE = 22). Table 2 shows the protective effects of polyols against pressure deactivation at 2500 bar. From these observations, one can derive for ~-galactosidase the following ranking of stabilisation efficiency of the four polyols at 2M: glycerol (C3) < sorbitol (C6), erythritol (C4) < xylitol (C5). Protective effects obtained with polyols are particularly high compared with those obtained in the presence of salts, even with the most stabilising one, n~mely KC1.
208 25 ..
20
15-
0
10-"
o
1.
2
50
I
0
_. ........... -~........................ ,,,--___-..... T T 1 2 [polyols] (M)
Figure 2. Protective effect of polyols on ~-galactosidase at 50~ as a function of their concentration: glycerol ( e e ), erythritol (" -), xylitol ( ~ ~ ) and sorbitol (~--o). Table 2. Protective effect of polyols on ~-~alactosidase at 2500 bar [Poly01s ] (M)
Glycerol
Erythritol
Xylitol
Sorbitol
0
1_+0.1
1_+0.1
1_+0.1
1_-20.1
0.5
5_+0.6
175+44
22+4
0.6_+0.2
1
14+1
278+_44
75+_36
40+_8
2
600+_56
4550_+900
11030+_1590
1625+_158
The weak effect of polyols on water structure [9] seems to indicate t h a t the protective ability of these additives during pressure denaturation can be connected to a direct interaction (specific or non-specific) between the polyhydric alcohol molecule and the protein. Our results show that enzyme stability may be very greatly increased by precise control of the nature and structure of its molecular micro-environment. This control is here achieved through the addition of specific compounds in the enzymatic solution. More information about the effects of soluble additives on enzyme barostability may be found in an article by Athes and Combes [10].
3. I N F L U E N C E O F P R E S S U R E ON T H E R M A L I N A C T I V A T I O N Parallel to the possibility of denaturing proteins, a stabilising effect of high pressure against thermal inactivation has been reported for several enzymes [4], so the influence of pressure on the thermal inactivation kinetics at
209 45~ of ~-galactosidase was investigated. To estimate the stabi]ising effect of pressure against thermal inactivation at moderate pressures, half-lives have been determined as a function of pressure and are reported in Figure 3. The observed stabilisation appeared to be maximal at 100 MPa, where the half-life at 45~ was 36 rain whereas it was only 15 rain at atmospheric pressure and 11 min at 150 MPa. 40 30.~ 2 0 -
/
10
0
I
I
I
I
1
50
100
150
200
250
300
Pressure (MPa) Figure 3. Influence of pressure on the half-life times at 45~ ~-galactosidase.
of K. lactis
Table 3 shows the remaining activities of E. coli and A. oryzae ~galactosidases solutions, which have been determined in standard conditions after a one hour incubation period at 55~ at pressures ranging from atmospheric pressure to 400 MPa. Since K. lactis ~-galactosidase is very sensitive to temperature (tin = 1.3 rain under atmospheric pressure at 55~ the effect of pressure on the thermal stability of this enzyme has been investigated at 45~ (tin = 15 rain under atmospheric pressure at 45~ Table 3. Residual activities (expressed in percent of the initial enzymatic activity) for a 1 hour thermal treatment at different pressures, for K. lactis ~-~alactosidase at 45~ and E. coli and A. or~/zae ~-~alactosidases at 55~ Pressure (MPa)
0.1
50
100
150
200
300
400
45~ / K. lactis ~-galactosidase
13%
63%
84%
74%
19%
0%
0%
55~ /A. oryzae ~-galactosidase
0%
51%
85%
92%
96%
93%
69%
55~
60%
79%
82%
81%
79%
70%
10%
/ E. coli ~-galactosidase
the 100% is obtained for native enzymes
210 Application of high hydrostatic pressure in the range 50-300 MPa for E. coli ~-galactosidase, 50-400 MPa for A. oryzae ~-galactosidase and 50-150 MPa for K. lactis ~-galactosidase stabilized these mesophilic enzymes against thermal inactivation. The respective pressure ranges of stabilization against thermal denaturation for each enzyme correspond to the range in which pressure alone does not lead to a significant inactivation [5]. 4. CONCLUSIONS For K. lactis ~-galactosidase, pressure and/or the use of some soluble additives, namely KC1 and polyols, could then be used to increase noticeably its thermal stability; this could be of great biotechnological importance when carrying out enzymatic conversions at high temperatures. Moreover, the use of polyols to increase the enzyme barostability could also be of great interest to carry out new ~-galactosidase-catalysed reactions at high pressures. REFERENCES
1. J.C. Cheftel, Food Sci. Technol. Int., 1 (1995) 75. 2. C. Balny, P. Masson and F. Travers, High Pres. Res., 2 (1989) 1. 3- D. CavaiUe and D. Combes, J. Biotechnol., 43 (1995) 221. 4. V.V. Mozhaev, R. Lange, E.V. Kudryashova and C. Balny, Biotechnol. Bioeng., 52 (1996).320. 5. V. AthOs, P. Degraeve, D. Cavaill4-Lefebvre, S. Espeillac, P. Lemay and D. Combes, Biotech. Letters, 19 (1997) 273. 6. P.C. Michels, D. Hei and D.S. Clark, Adv. Prot. Chem., 48 (1996) 341. 7. R.V. Rariy, N. Bec, N.L. Klyachko, A.V. Levashov and C. Balny, Biotechnol. Bioeng., 57 (1998) 552. 8. R.D., Schmid, Adv. Biochem. Eng., 12 (1979) 41. 9. I. Auzanneau, D. Combes and A. Zwick, J. R~man Spectrosc., 22 (1991) 227. 10. V. AthOs and D. Combes, Enzyme Microb. Technol., 22 (1998) in press.
Stability and Stabilization of Biocatalysts A. BaUesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
211
T h e r m o d y n a m i c stability of a m o n o m e r i c derivative of bovine seminal ribonuclease G. Barone a, F. Catanzano a, G. Graziano b, V. Cafaro c, G. D'Alessio c and A. Di Donato c aDept. Chemistry, University "Federico II", Via Mezzocannone, 4 - 80134 Naples, Italy bFaculty of Science, University of Sannio, Via Marmorale- 82020 Paduli (BN), Italy CDept. Biochem., University "Federico II", Via Mezzocannone, 16- 80134 Naples, Italy
Bovine seminal ribonuclease, BS-RNase, is a dimeric enzyme possessing very special biological actions. Its identical subunits possess more than 80 % of sequence identity with pancreatic RNase A. We prepared a monomeric and carboxyamidomethylated form of bovine seminal ribonuclease, MCAM-BS-RNase, in order to compare its thermodynamic stability with that of RNase A. DSC and CD measurements indicated that MCAM-BS-RNase has a reduced thermal stability with respect to RNase A.
I. INTRODUCTION Bovine seminal ribonuclease, BS-RNase, is the only dimeric ribonuclease of the pancreatic-type superfamily discovered so far [ 1]. Its two identical subunits are linked by two disulfide bridges pairing Cys31 and 32 of one chain with Cys32 and 31, respectively, of the other chain, and possess more than 80 % of sequence identity with pancreatic ribonuclease, RNase A. The two subunits are able to assume two quaternary structures [2]: the MxM form in which there is the swapping of the N-terminal segment between the two subunits [3], and the M=M form in which the swapping does not occur and the N-terminal segment interacts with the remaining part of its own subunit [2]. The two quaternary structures are in equilibrium between each other in the natural protein isolated from bull seminal plasma in a molar ratio [MxM]/[M=M] ~, 2/1 at physiological temperature [2]. To this extraordinary property, it is necessary to add that BS-RNase possesses special biological actions: for instance, the MxM form is a cytotoxic agent selective for some tumoral cells [4]. Selective cleavage of the intersubunit disulfide bridges with a moderate excess of dithiotreitol, followed by alkylation of the exposed sulfydril groups with iodoacetamide, allows the preparation of carboxyamidomethylated monomers of BS-RNase [5]. This monomeric derivative is folded, stable and enzymatically active, but is devoid of all the special biological actions of dimeric BS-RNase. We considered interesting to compare the
212 thermal stability of MCAM-BS-RNase and RNase A in view of the high sequence identity: 101 out of 124 residues are identical; the eight Cys residues, that pair to form the four intrachain disulfide bridges, occur at identical sequence positions; the residues that make up the catalytic site are identical. We performed differential scanning calorimetry, DSC, and circular dichroism, CD, measurements at pH 5.0. The data pointed out that MCAM-BS-RNase is less thermally stable than RNase A. On the basis of the structural information available for RNase A and dimeric and monomeric forms of BS-RNase, an explanation is proposed.
2. MATERIALS AND METHODS
Carboxyamidomethylated monomers of BS-RNase were obtained as described [5], while RNase A was type XII A of SIGMA. Protein purity was checked by SDS-PAGE, whereas protein concentration was calculated spectrophotometrically by using A(0.1%, 278 nm) = 0.71 for RNase A and 0.54 for MCAM-BS-RNase [6]. CD spectra in the near and farUV region were recorded with a Jasco J-710 spectropolarimeter, as described elsewhere [7]. DSC measurements were carried out on a second-generation Setaram Micro-DSC apparatus, interfaced with a data translation A/D board for automatic data accumulation [7]. A scan rate of 0.5 K min -l was chosen for the present study. The excess heat capacity function
was obtained after baseline subtraction, assuming that the baseline is given by the linear temperature dependence of native state heat capacity. The calorimetric enthalpy AdH(Td) was determined by direct integration of the area under the curve, and the van't Hoff enthalpy was calculated with the standard formula: AdHv.H.(Td) = 4 RTd2 [/AdH(Td) ]
(i)
where T d is the denaturation temperature and corresponds to the maximum of the DSC peak, is the value of the excess molar heat capacity function at T d, and R is the gas constant. The closeness to 1 of the cooperative unit, defined as the calorimetric to van't Hoff enthalpy ratio, CU - AdH(Td)/AdHv.H.(Td) , is a necessary condition to state that the denaturation is a two-state transition.
3. RESULTS AND DISCUSSION DSC measurements were performed at pH 5.0, 100 mM acetate buffer, to compare the thermal stability of MCAM-BS-RNase with that of RNase A. The results are collected in Table 1 and representative DSC curves are shown in Figure 1. The temperature-induced transitions of these proteins were reversible, not influenced by protein concentration (in the range 1.4 - 3.0 mg mL-l), and well represented by the two-state N <::>D transition model, as
213
Table 1. Thermodynamic parameters of the temperature-induced denaturation of MCAM-BS-RNase and RNase A at pH 5.0, obtained from DSC and CD measurements.
MCAM-BS-RNase
RNase A
Td (~
AdH(Td) (kJmo1-1)
AdS(Td) (kJ K-lmo1-1)
AdCp (kJ K-lmol "1)
CD222
55.1
375 + 15
1.14 _+0.05
. . . .
CD275 DSC
55.0
370 + 16
1.13 + 0.05
. . . .
55.2
380 + 12
1.16 + 0.04
4.7 + 0.5
CD222
61.2
460 _ 14
1.38 _+0.05
--
CD275
61.1
455 +_ 15
1.36 _ 0.05
--
DSC
61.3
465 _+ 13
1.39 _+0.04
5.5 _+0.6
CU
0.97
1.01
Note. Each figure is the mean value of four measurements. The error in T d does not exceed 0.2 ~ Reported errors for AdH(Td) and AdC p are the standard deviations of the mean from the multiple determinations. Reported errors for AdS(Td) are calculated by propagating the errors for AdH(Td) and T d.
60
b
(kJ K-lmo1-1)
i
40
I
I
temperature (~
i
70
Figure 1. DSC profiles at pH 5.0, 100 mM acetate buffer, of MCAM-BS-RNase (curve a) and RNase A (curve b).
214 emphasized by the closeness to 1 of the cooperative unit. MCAM-BS-RNase showed a denaturation temperature of 55.2 ~ and an enthalpy change of 380 kJ mol-l.These values are 6.1 ~ and 85 kJ mo1-1, respectively, lower than the corresponding ones of RNase A. This demonstrates that, although MCAM-BS-RNase does possess more than 80 % of sequence identity with RNase A, it is markedly less stable toward temperature. In addition, AdCp = 4.7 + 0.5 kJ K-lmo1-1 for MCAM-BS-RNase, and 5.5 + 0.6 kJ K-lmol -l for RNase A. The lowering in AdCp should be due to a greater exposure to water of nonpolar groups in the folded conformation, in the assumption that the denatured state is identical for the two proteins [8]. Actually, the thermal stability of the monomeric form of BSRNase is not affected by the nature of the alkylating agent used to protect the free sulfydril groups. In fact, a monomeric and carboxymethylated derivative, MCM-BS-RNase, showed T d = 55.8 ~ and AdH(Td) = 365 kJ mo1-1, while a BS-RNase monomer linked through mixed disulfides with two glutathione moieties showed Td = 55.4 ~ and AdH(Td) = 375 kJ mo1-1, always at pH 5.0, 100 mM acetate buffer. CD measurements in the far-UV region showed that the content of secondary structure of the two proteins is practically identical, whereas some differences were recorded in the near-UV region. In fact, the intensity of the negative band centered at 275 nm, characteristic of RNase A and caused by the presence of six tyrosine residues (i.e., Tyr25, 73, 76, 92, 97, and 115), is reduced for MCAM-BS-RNase because it possesses only four tyrosine residues (i.e., it lacks Tyr76 and 115). The thermal denaturation profiles, obtained by monitoring the CD signal both at 222 nm and 275 nm, pH 5.0, 10 mM acetate buffer, resulted reversible, and were analyzed by van't Hoff procedure, assuming valid the two-state N r D model, as demonstrated by DSC measurements. The values of Td, AdH(Td) and AdS(Td) are in line with those coming from DSC scans, as can be seen in Table 1. Similar results were obtained by means of CD and fluorescence measurements for MCM-BS-RNase [9]. A correct thermodynamic comparison requires the evaluation of AdH and AdS at the same temperature for the two proteins. We selected, as reference, the T d of MCAM-BSRNase, 55.2 ~ The calculations for RNase A gave AdH(55.2 ~ = 431 + 13 kJ tool -1 and ~dS(55.2 ~ = 1.29 + 0.04 kJ K-lmol -!. These values were 51 kJ tool -1 and 130 J K-lmo1-1, respectively, greater than the corresponding ones of MCAM-BS-RNase. Therefore, the destabilization of the monomeric derivative of BS-RNase with respect to the pancreatic enzyme was due to enthalpic factors, only partially counterbalanced by entropic factors. It is possible to correlate the thermodynamic results to structural data. The secondary structure in solution at pH 5.0 of MCM-BS-RNase has been recently determined by NMR [10]: it is practically identical to that of RNase A and each subunit of MxM-BS-RNase, as determined from X-ray crystallography [3]. Additionally, from the NMR data collected, an initial model of the tertiary structure of MCM-BS-RNase was constructed [10], and it has a folding pattern very similar to that of RNase A. However, some differences exist in the segments of the polypeptide chain containing several substitutions. The three-dimensional structure of RNase A [11 ] can be subdivided in a tail (residues 1-15) and a body (residues 23-124), linked by an irregularly folded heptapeptide (residues 16-
215 22), which is called hinge-peptide. Four amino acid substitutions occur in the hinge-peptide between the pancreatic and seminal enzymes, the most important of which is the replacement of Alal9 by Pro. Indeed, a trans conformation was assigned to Pro l9 in MCM-BS-RNase [ 10], which causes a shift toward the exterior not only of the hinge-peptide, but of the entire N-terminal segment. The authors stressed that the pattern of NOE signals observed between the residues of the N-terminal segment and those belonging to the body of the molecule brings out that the two parts do not interact as strongly as in RNase A. In addition, theoretical conformational energy calculations, carried out on the hinge-peptide of RNase A with the Alal9Pro substitution, predicted that Pro l9 assumes a trans conformation causing a large conformational change of the hinge-peptide, which is shifted toward the exterior of the molecule [12]. These findings explain the large conformational freedom of the hinge-peptide of BS-RNase, which is able to assume two different conformations in the MxM and M=M quaternary forms [2]. The structural evidence arising from NMR data and theoretical calculations is in agreement with the substantial difference in thermal stability existing between RNase A and MCAM-BS-RNase. In addition, we have shown in a previous study that 4 out of the 23 amino acid substitutions between RNase A and MCAM-BS-RNase are mainly responsible for the decrease in thermal stability. In fact, the progressive introduction of a Pro, a Leu and two Cys at positions 19, 28, 31 and 32, respectively, caused a stepwise decrease in thermal stability of RNase A, gradually transforming it into a virtual monomer of BS-RNase [ 13,14]. Finally, it was found that BS-RNase, the equilibrium mixture of MxM and M=M, has T d = 61.6 ~ at pH 5.0 [ 14], a value similar to that of RNase A, but greater than that of its monomeric derivative. This finding can be rationalized on structural grounds. The two intersubunit disulfide bridges give rise to a sixteen membered ring encompassing four residues in BS-RNase. The presence of two covalent linkages between the two subunits should stabilize the protein by reducing the conformational entropy of the denatured state relative to the native state. It has been proposed [ 15] that the observed changes in stability of a number of proteins can be rationalized by the following equation: ASconf = - 14.4 - 1.5.R.InN
(2)
where ASconf is the decrease in conformational entropy of a randomly coiled polypeptide chain caused by the introduction of a disulfide bridge fixing N residues, and R is the gas constant, 8.314 J K-lmo1-1. By applying this equation to dimeric BS-RNase with N = 4, one obtains ASconf = -32 J K-lmol -l . Each subunit would be stabilized for a quantity equal to half of the calculated value, -16 J K-lmo1-1, because only two residues for each subunit are fixed by the two disulfide bridges. As a consequence, the denaturation temperature of MCAM-BSRNase would become: Td'= AdH(Td)/[AdS(Td) - ASconf] = 380 (kJ mo1-1)/(1.157 - 0.016) (kJ K-lmol -l) = = 333.05 K = 59.9 ~
(3)
216 which in slightly lower than the experimental value of 61.6 ~ In order to reach a complete agreement one should take into account that the two subunits, beyond the two covalent links, pack close together, on dimer formation, through a-helices encompassing residues 24-34, whose axes form an angle of about 40 degrees [3]. Such an interface provides additional stability to the dimeric structure by allowing good van der Waals interactions between the side-chains of Leu28 and Met29 of both subunits. In conclusion, the two interchain disulfide bridges provide a rigid scaffold for the subunit interface and render the dimeric structure of BS-RNase as stable toward temperature as RNase A, although its individual subunits possess a lower thermal stability.
Acknowledgments. Work supported by P.R.I.N. grant from the Italian Ministry for University and Scientific and Technological Research (M.U.R.S.T., Rome) and University of Naples "Federico Ir'.
REFERENCES
1. G. D'Alessio, A. Di Donato, L. Mazzarella and R. Piccoli, in Ribonucleases: Structures and Function (G. D'Alessio and J.F. Riordan, Eds.) pp 383-423, Academic Press, New York, 1997. 2. R. Piccoli, M. Tamburrini, G. Piccialli, A. Di Donato, A. Parente and G. D'Alessio, Proc.Natl.Acad.Sci. U.S.A., 89 (1992) 1870. 3. L. Mazzarella, S. Capasso, D. Demasi, G. Di Lorenzo, C.A. Mattia and A. Zagari, Acta Cryst., D49 (1993) 389. 4. R.J. Youle and G. D'Alessio, in Ribonucleases: Structures and Function (G. D'Alessio and J.F. Riordan, Eds.) pp 491-514, Academic Press, New York, 1997. 5. G. D'Alessio, M.G. Malomi and A. Parente, Biochemistry, 14 (1975) 1116. 6. A. Parente, D. Albanesi, A.M. Garzillo and G. D'Alessio, Ital. J.Biochem., 26 (1977) 451. 7. F. Catanzano, C. Giancola, G. Graziano and G. Barone, Biochemistry, 35 (1996) 13378. 8. G. Graziano, F. Catanzano, P. Del Vecchio, C. Giancola and G. Barone, Gazz.Chim.It., 126 (1996) 559. 9. C. Grandi, G. D'Alessio and A. Fontana, Biochemistry, 18 (1979) 3413. 10. A. D'Ursi, H. Oschkinat, C. Cieslar, D. Picone, G. D'Alessio, P. Amodeo and P.A. Temussi, Eur.J.Biochem., 229 (1995) 494. 11. A. Wlodawer, L.A. Svensson, L. Siolin and G.L. Gilliland, Biochemistry, 27 (1988) 2705. 12. L. MazzareUa, L. Vitagliano and A. Zagari, Proc.Natl.Acad.Sci. U.S.A., 92 (1995) 3799. 13. A. Di Donato, V. Cafaro, I. Romeo and G. D'Alessio, Protein Sci., 4 (1995) 1470. 14. F. Catanzano, G. Graziano, V. Cafaro, G. D'Alessio, A. Di Donato and G. Barone, Biochemistry, 36 (1997) 14403. 15. P.M. Harrison and M.J.E. Sternberg, J.Mol.Biol., 244 (1994) 448.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
T h e r m o d y n a m i c stability o f r i b o n u c l e a s e P2 from and s o m e m u t a n t s
217
Sulfolobus solfataricus
G. Graziano a, F. Catanzano b, P. Fusi c, P. Tortora c and G. Barone b aFaculty of Science, University of Sannio, Via Marmorale - 82020 Paduli (BN), Italy bDept. Chemistry, University "Federico II", Via Mezzocannone, 4 - 80134 Naples, Italy CDept. Biochemistry, University of Milan, Via Celoria, 26 - 20133 Milan, Italy
Ribonuclease P2 from the thermoacidophilic archaebacterium Sulfolobus solfataricus is a small globular protein with a known three-dimensional structure. Inspection of the structure reveals that Phe31 is a member of the aromatic cluster forming the protein hydrophobic core, whereas Trp23 is located on the protein surface and its side-chain is exposed to the solvent. The thermodynamic consequences of the substitution of these two residues in ribonuclease P2 have been investigated by comparing the temperature-induced denaturation of P2 with that of three mutants: F31A-P2, F31 Y-P2 and W23A-P2.
1. INTRODUCTION Ribonuclease P2 is a small basic protein recently isolated from Sulfolobus solfataricus [1], a thermoacidophilic archaebacterium that lives at 87 ~ and acidic pH in volcanic hot springs [2]. P2 is a 62 residue protein with no histidine or asparagine, and no cysteine; its encoding gene was cloned and expressed in E. coil [3]. The recombinant protein was identical to the natural form, the only difference being the absence of monomethylation at Lys4 and Lys6. P2, apart from its ribonuclease activity [1], it acts as a non-specific DNA-binding protein, protecting DNA from thermal denaturation [4]. The NMR solution structure [5,6], shows that the protein has a compact folding pattern composed of a three-stranded 13-sheet orthogonal to a two-stranded fl-sheet, and with only a small helical stretch at the C-terminus. Actually, Hard's group solved the structure of Sso7d, which differs from P2 for the presence of another lysine residue at the C-end. Therefore, the structure and stability of P2 can safely be considered identical to those of Sso7d. P2 has a hydrophilic surface with many basic residues (i.e., 2 arginines and 13 lysines), and a very hydrophobic core with a cluster of aromatic sidechains (Phe5, Tyr7, Phe31 and Tyr33) in herringbone geometry. Its small size allowed the construction and expression of several mutant forms. Therefore, P2 is an ideal candidate to try to clarify the molecular origin of the extra-stability of thermophilic proteins. A comprehensive
218 study on the thermal stability of P2 by Ladenstein and co-workers [7] pointed out that the protein has a denaturation temperature of 98.2 ~ in the pH range 4.5-7.0 and the process is a reversible two-state N r D transition. Thermodynamic characterization of three mutants of P2: F31A-P2, F31Y-P2 and W23A-P2 is the subject of this work. The sites of amino acid substitutions were suitably selected in order to bring out the structural determinants of protein thermal stability. Indeed, Phe31 is a member of the aromatic cluster considered fundamental for the stability of native conformation. In contrast, Trp23 is located on the protein surface and its side-chain seems to play a major role in the DNA binding process [4]. We carried out DSC measurements in a wide range of pH: all the three mutants proved to be less thermally stable than the parent protein. These findings are discussed and a rationale is proposed.
2. MATERIALS AND METHODS
P2 and its mutants were produced as previously reported [8]. Protein concentration was determined spectrophotometrically using extinction coefficients calculated from the absorption of tyrosine, e280 = 1400 M-lcm -l, and tryptophan, e280 = 5500 M-lcm -l [9]. DSC measurements were carded out as described elsewhere [ 10]. The overall molar heat effect of denaturation is not large because of the small size of P2, and to get reliable records of the heat capacity profiles, we had to work at relatively high protein concentration (2.5-5.5 mg mL-1); nevertheless, neither concentration nor heating rate dependence (in the range 0.5-1.5 K min -l) of the heat capacity peaks was observed. Moreover, the DSC peaks extend beyond 100 ~ in most of the experiments; as the operational range of our calorimeter is limited to 100 ~ incomplete DSC peaks were recorded, and a different analysis from the conventional one was carded out [ 10]. The excess molar heat capacity function for a two-state N r D transition is: = [AdH(Td) + AdCp(T-Td)].(dfD/dT ) + fD.AdCp
(1)
where fD = [Kd/(l+Kd)] is the fraction of denatured molecules, K d is the corresponding equilibrium constant, AdH(Td) and AdC p represent the denaturation enthalpy and heat capacity changes. A nonlinear regression of the experimental DSC profiles with respect to Eq.(1) was performed, obtaining estimates for Td, AdH(Td) and AdCp. A close agreement between the calculated and the experimental profile has to be considered as a necessary and sufficient condition to conclude that the denaturation is a two-state N r D transition.
3. RESULTS AND DISCUSSION
DSC measurements were performed in 10 mM buffer with 100 mM NaC1, using a scan rate of 1 K min -l in the pH range 2.1-7.2. In these experimental conditions, the thermal
219 denaturation of the proteins investigated was a reversible two-state transition. A series of DSC profiles for F3 IA-P2 and F31 Y-P2 are shown in Figures 1 and 2.
10 (kJ K'lmo1-1)
20
temperature (*C)
90
Figure 1. DSC peaks for F31A-P2 at different pH values: curve a at pH 3.0; curve b at pH 3.5; curve c at pH 6.0. The continuous curves through the experimental points are the results of the nonlinear regression with respect to Eq.(1).
15
•
(kJ K'I tool "1)
30
temperature (°C)
90
Figure 2. DSC peaks for F31 Y-P2 at different pH values: curve a at pH 2.6; curve b at pH 3.0; curve c at pH 4.0; curve d at pH 5.5. The continuous curves through the experimental points are the results of the nonlinear regression with respect to Eq.(1).
220 The T d values of P2 and its three mutants showed a marked pH dependence up to pH around 5.0, and then practically leveled off, remaining constant up to pH 7.0. All the three mutants are less thermally stable than the parent protein; in the pH range 5.0-7.0, the F31A substitution leads to a decrease of 24 ~ in the T d value, the F31Y substitution to a decrease of 10 ~ and the W23A substitution to a decrease of 6 ~ On the other hand, the values of AdC p, calculated for P2 and its three mutants from the AdH(Td) vs T d plots, are very similar among each other, amounting to about 2.6 kJ K-lmol -l . On the basis of such findings, we performed a comparison of the thermodynamic stability of P2 with that of its three mutants. The mean values of T d, AdH(Td), AdS(T d) and AdC p in the pH range 5.0-7.0 for the four proteins are collected in Table 1.
Table 1.
Average values of thermodynamic parameters for ribonuclease P2 and its three mutants in the conditions of maximal thermal stability. Data from ref. [ 10]. protein
Td (~
AdH(Td)
AdS(Td)
AdH(98.2 ~
(kJ mo1-1) (J K-lmo1-1)
ADS(98.2 ~
AdC p
(kJ mol" I)
(J K-lmo1-1)
(kJ K-lmol -!)
,
P2
98.2
267 + 5
719 + 14
267 + 5
719 + 14
2.6 + 0.3
F31A-P2
74.4
186 + 5
535 + 15
246 + 7
701 + 20
2.5 + 0.2
F31 Y-P2
88.0
223 + 4
618 + 11
250 + 5
691 + 14
2.6 + 0.3
W23A-P2
92.2
247 + 2
676 + 6
263 + 3
720 + 7
2.7 + 0.2
These thermodynamic parameters allow the determination of the stability curves [ 11 ], AdG vs T, of the four proteins, according to the equation: AdG(T) = AdH(Td)[1 - (T/Td)] + AdCp[T - T d - T ln(T/Td) ]
(2)
which is exact in the assumption that AdC p is temperature-independent. A comparison among the calculated stability curves indicates that the amino acid substitutions do not practically affect the point of maximal stability, as it occurs always around 285 K. However, the corresponding values of AdG are different: 33.7 kJ mo1-1 for P2, 28.5 kJ mol -I for W23A-P2, 24.5 kJ mo1-1 for F31Y-P2, and 18.6 kJ mop ! for F31A-P2. The maximum value of AdG for P2, when normalized per residue, amounts to 540 J mo1-1, a value comparable to those determined for globular proteins isolated from mesophilic organisms [12]. This finding validates the suggestion by Jaenicke I13] that the stability curves of thermophilic proteins are flattened with respect to those of mesophilic counterparts.
221 In view of the low curvature of the stability curves, the values of AdG at 25 ~ are very close to those corresponding to the point of maximum. Therefore, AAdG(25 ~ = 15 kJ mol "! for F31A-P2, AAdG(25 ~ = 9 kJ mo1-1 for F31Y-P2, and AAdG(25 ~ = 5 kJ mol "l for W23A-P2. These experimental values can be compared to those calculated on the basis of the difference in side-chain hydrophobicity. According to the "pure" hydrophobicity scale recently proposed [14], AAdG(25 ~ = 5.4 kJ mo1-1 for a mutation Phe---~Ala at a buried position, AAdG(25 ~ = 2.9 kJ mol -l for a mutation Phe---~Tyr, AAdG(25 ~ 9.0 kJ mol "! for a mutation Trp~Ala. The discrepancy in the case of W23A-P2 is in part due to the fact that the side-chain of residue 23 is not completely buried. However, in general, the values of AAdG(25 ~ calculated on the basis of hydrophobicity alone do not quantitatively agree with the experimental ones. This seems to suggest that other factors, beyond the hydrophobic effect, are important in determining the stability of folded conformation. In order to reach a deeper understanding of the molecular origin of the difference in thermodynamic stability, we calculated AdH and AdS at a common temperature for the four proteins, selecting, as reference, the T d of P2, 98.2 ~ to minimize extrapolation errors. The values of AdH(98.2 ~ and ADS(98.2 ~ for P2, F31A-P2, F31Y-P2 and W23A-P2 are collected in the fifth and sixth columns of Table 1. The F3 I A and F31Y substitutions affect only slightly the entropy change, but cause a decrease in the average value of AdH(98.2 ~ from 267 + 5 kJ mo1-1 to 246 + 7 kJ mo1-1 and 250 9 5 kJ mo1-1, respectively. This suggests that the destabilization of folded structure is of enthalpic origin. The F31A substitution causes a loss of van der Waals interactions in the protein hydrophobic core, as the volume of Phe side-chain is much larger than that of Ala side-chain, 129.7 A 3 against 26.3 .&3 [15]. In addition, the partial destruction of the aromatic cluster Phe5, Tyr7, Phe31 and Tyr33 should cause an enthalpic destabilization of native structure [16]. In fact, molecular dynamics calculations brought out that Phe31 has the largest van der Waals energy among all the amino acid residues of P2 [ 17], because Phe31 is located in a strongly nonpolar region constituted by the side-chains of Val3, Vall4, Ile19, Va122, Ile29, Va145 and Leu54, beyond the already mentioned aromatic cluster. In the case of the F31Y substitution, the aromatic cluster should remain intact and no cavity should be created, as the side-chain volume is 129.7 ,&3 for Phe and 133.3 A 3 for Tyr [15]. Therefore, the enthalpic penalty associated with the dehydration and burial in the protein hydrophobic core of the hydroxyl group should be the cause of the destabilization [18]: they should not be counterbalanced by the formation of an intramolecular hydrogen bond because the region is very rich in nonpolar moieties. The W23A substitution has no effect on the entropy change and causes a very slight decrease in the average value of AdH(98.2 ~ 263 a: 2 kJ mol -! vs 267 + 5 kJ mo1-1. The large aromatic side-chain of Trp is exposed to the solvent and should interact more favourably with water than the small aliphatic side-chain of Ala, through the formation of hydrogen bonds involving the delocalized n electrons of the ring [19]. In addition, the favourable van der Waals interactions existing between the large side-chain of Trp23 and other groups of the protein [17] are probably lost due to the smallness of Ala side-chain.
222 In conclusion, DSC data indicate that the hydrophobic core is the main responsible of the great thermal stability of P2, even though other factors are important, as suggested by the F31Y and W23A substitutions.
Acknowledgment. Work supported by P.R.I.N. grant from the Italian Ministry for University and Scientific and Technological Research (M.U.R.S.T., Rome) and University of Naples "Federico II".
REFERENCES 1. P. Fusi, G. Tedeschi, A. Aliverti, S. Ronchi, P. Tortora and A. Guerritore, Eur.J.Biochem., 211 (1993) 305. 2. M. De Rosa, A. Gambacorta, B. Nicolaus, P. Giardina, E. Poerio and V. Buonocore, Biochem.J., 224 (1984) 407. 3. P. Fusi, M. Grisa, E. Mombelli, R. Consonni, P. Tortora and M. Vanoni, Gene, 154 (1995) 97. 4. H. Baumann, S. Knapp, A. Karshikoff, R. Ladenstein and T. Hard, J.Mol.Biol., 247 (1995) 840. 5. H. Baumann, S. Knapp, T. Lundback, R. Ladenstein and T. Hard, Nat.Struct.Biol., 1 (1994) 808. 6. R. Consonni, R. Limiroli, H. Molinari, P. Fusi, M. Grisa, M. Vanoni and P. Tortora, FEBS Lett., 372 (1995) 135. 7. S. Knapp, A. Karshikoff, K.D. Bemdt, P. Christova, B. Atanasov and R. Ladenstein, J.Mol.Biol., 264 (1996) 1132. 8. P. Fusi, K. Goossens, R. Consonni, M. Grisa, P. Puricelli, G. Vecchio, M. Vanoni, L. Zetta, K. Heremans and P. Tortora, Proteins, 29 (1997) 381. 9. S.C. Gill and P.H. von Hippel, Anal.Biochem., 182 (1989) 319. 10. F. Catanzano, G. Graziano, P. Fusi, P.Tortora, G. Barone, Biochemistry, accepted for publication. 11. W.J. Becktel and J.A. Schellman, Biopolymers, 26 (1987) 1862. 12. G.I. Makhatadze and P.L. Privalov, Adv.Protein Chem., 47 (1995) 307. 13. R. Jaenicke, Eur.J.Biochem., 202 (1991) 715. 14. P.A. Karplus, Protein Sci., 6 (1997) 1302. 15. Y. Harpaz, M. Gerstein and C. Chothia, Structure, 2 (1994) 641. 16. S.K. Burley and G.A. Petsko, Science, 229 (1985) 23. 17. E. Mombelli, M. Afshar, P. Fusi, M. Mariani, P. Tortora, J.P. Connelly and R. Lange, Biochemistry, 36 (1997) 8733. 18. A.S. Yang, K.A. Sharp and B. Honig, J.Mol.Biol., 227 (1992) 889. 19. P. Linse, J.Am.Chem.Soc., 112 (1990) 1744.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
223
Stabilization of ~ - C h y m o t r y p s i n by D M S O E. Flaschel and L. Ebmeier Universit&t Bielefeld, Technische Fakult&t, Postfach 10 01 31, D-33501 Bielefeid, Germany 1.
INTRODUCTION
o~-Chymotrypsin is a protease with proteolytic as well as esterolytic activity. Besides processes based on stereospecific hydrolysis, ~-chymotrypsin may be used for the synthesis of peptides (Kisee et aL 1988) and for transesterification (Moresoli et aL 1992a) under suitable reaction conditions. Due to autolysis o~-chymotrypsin is most often used in immobilized form. Depending on the reaction system, however, the effectiveness of immobilized enzymes may be low because of pore diffusion and other mass-transfer phenomena. Therefore, strategies are sought for stabilizing ~chymotrypsin in soluble form. Besides chemical modifications of the enzyme and the addition of salting-in or salting-out additives, an unusual stabilization of ~chymotrypsin with half-lives up to 200 days has been observed in the presence of 10 and 20 % (v/v) dimethyl sulfoxide (DMSO) (Pliura and Jones 1980; Moresoli et al. 1992b). Motivated by these observations the stability of o~-chymotrypsin was investigated in a larger domain of the most essential reaction variables. For this purpose, the DMSO concentration, the temperature as well as the pH were varied. 2.
MATERIALS AND METHODS
o~-Chymotrypsin (C-4129) and N-benzoyI-L-tyrosine ethyl ester (BTEE) (B-6125) were purchased from Sigma, dimethyl sulfoxide (DMSO) from J.T. Baker (7033), N~succinyl phenylalanine p-nitroanilide (SUPHEPA) from Merck (12444,0001), and azocasein from Fluka (11610). All other reagents were at least of analytical grade. For studying the influence of DMSO on the stability of o~-chymotrypsin the enzyme was stored in a Tris-maleate/NaOH buffer adjusted to the appropriate pH and containing 20 mmol/I CaCI2. The solution consisted of 0.2 M Tris as well as 0.2 M maleic acid. The enzyme solutions prepared with the appropiate concentration of DMSO were stored in a dry thermostat (Kleinfeld BT 100) by means of 4 ml test tubes at the appropriate temperature. The activity of cz-chymotrypsin was monitored periodically by means of the SUPHEPA test.
224 The SUPHEPA test was based on the hydrolysis of N%succinyl phenylalanine pnitroanilide (SUPHEPA) by spectrophotometry in thermostated cuvettes at 25 ~ and a wavelength of 405 nm (~4o5= 1.02 I/(mmol mm)). A SUPHEPA solution was prepared with a triethanol amine (TEA) buffer containing 0.2 M TEA adjusted to pH 7.8 by NaOH. The concentration of SUPHEPA was 12 mmol/l. A volume of 0.95 ml of another triethanol amine buffer containing in addition 20 mmol/I of CaCI2 was transferred into a cuvette together with 50 IJI of the o~-chymotrypsin solution to be tested. After 5 min of temperature equilibration, the assay was started by adding 0.5 ml of the SUPHEPA solution. The increase of absorption was monitored for about 30 min. In the case of studies of the activity of o~-chymotrypsin in the presence of DMSO the appropriate concentration of DMSO was supplied with the TEA buffer. The BTEE test was based on the fact that the product of the esterolytic action, NbenzoyI-L-tyrosine (BT) absorbes light at a wavelength of 250-260 nm (~256 = 964 I/(mol cm)). The test was performed in thermostated quartz cuvettes at 25 ~ BenzoyI-L-tyrosine ethyl ester (BTEE) was solubilized in DMSO in order to give a 15.5 mM solution. A volume of 3 ml of a Tris-HCI buffer prepared with the appropriate concentration of DMSO was thermostated in a cuvette for at least 5 min. A volume of 100 pl of the BTEE solution was added and the contents of the cuvette was mixed. The activity measurement was started by adding 100 pl of a solution of o~-chymotrypsin. After mixing the absorption was monitored for about 5 min. For the azocasein activity test a solution of 25 g/I was prepared by solubilizing azocasein in a 0.2 M Na2HPO4 solution, adjusting the pH by a 0.2 M NaH2PO, solution to 8.3 and filling up with water to twice the initial volume of phosphate buffer. In addition, an ~-chymotrypsin solution of 50 mg/I was prepared in a 0.1 M sodium phosphate buffer and the appropriate amount of DMSO. A volume of 200 IJl was equilibrated at 38 ~ in a test tube prior to adding 200 pl of the enzyme-DMSO solution. After mixing it was incubated for 35 min at 38 ~ A volume of 1.6 ml trichloroacetic acid was added, precipitating protein was eliminated by centrifugation for 15 min at 20,000 rpm. From the supernatent 1 ml was mixed with the same volume of a 0.5 M NaOH solution prior to determining the absorption at 450 nm. The deactivation of ~-chymotrypsin was modeled by a kinetics describing a firstorder consecutive reaction with a single intermediate:
kl
E
~ El
k2
~-- E2
(1)
The enzyme species E was the normal active enzyme species. E~ was a stable
enzyme species, whose concentration depended on the deactivation rate constants kl and k2 describing the consecutive reaction. The enzyme species E2 was supposed to be completely inactive, whereas the activity of the intermediate species E1 may show any ratio of activity with respect to the initial ~-chymotrypsin species E - defined by a factor 0~. According to this model, the residual relative activity - assuming that there was only the species E at the beginning - should evolve as:
a =
A = I 1 + ~ kl lexp(-k,t)a Ao k2 - k I
exp(-k2t ) ~ kl a k2 - k 1
For deactivation at 50 ~ the following simple first-order model has been applied:
(2)
225
a =
A
Ao
-
(3)
exp(-k2t )
assuming that the second step of deactivation became rate limiting. The activity ratio ot and the deactivation rate constants ka and k2 were obtained by means of the parameter-estimation method after Marquardt.
3.
RESULTS AND DISCUSSION
At 50 ~ and pH 6.0 DMSO has a destabilizing effect on ~-chymotrypsin as shown in Fig. 1. .4
...... Parameter:
~ 12 f >
DMSO
10
o 08 o ~ ,
>
concentration,
Co.so 1% (v/v) [] o v 30 L~ .to O 40 0 20
0.6 o 20
04
3O
0.240
00
0
9
I
15
~
I
,
I
30 45 Time, min
,
I
60
~
I
75
Figure 1" Deactivation of ot-chymotrypsin at 50 ~ and pH 6.0 When the temperature was lowered to 39 and 30 ~ just the opposite effect was observed: with increasing DMSO concentration o~-chymotrypsin was stabilized against thermal deactivation - as to be seen particularly well in Fig. 2. o~-Chymotrypsin solutions stored at temperatures of less than 44 ~ exhibited a remarkable increase of activity with time. In the absence of DMSO this activity increase during the initial deactivation phase was only observed for a maximum temperature of 39 ~ (data not shown). The degree of activation increased by lowering the temperature to 30 ~ By adding 10 % (v/v) DMSO this phenomenon was also observed at 44 ~ Higher DMSO concentrations caused the activity to stay constant at its maximum value for a longer time. However, the activity maximum flattened with increasing DMSO concentration. An increase of pH from 6.0 to 7.5 and 8.5 had a drastic effect on the stability of otchymotrypsin in the absence of DMSO - as shown in Fig. 3 for experiments at pH 7.5. Thus, the activity at pH 7.5 had dropped by 6 1 % after six days, whereas the activity at pH 6 had only dropped by 19 % after the same period of time.
226
1.4 3O
1.2 1.0 == 0.8 D
D
Parameter: couso !'1
(9
L.
0.4
0 V
m
<>
IZ
O
1% (vlv) 0
Z~ 10
-~ 0.2 , O
0
concentration,
DMSO
0.6
I
,
20 30 40 I
0
.
I
1
9
2
I
9
3
I
9
4
l
,
I
5
9
I
6
.
7
8
Time, days Figure 2" Deactivation of o~-chymotrypsin at 30 ~ and pH 6.0 1.4 30 20
1.2
40
Z~
10
0.8 t~ :3
"o o,m
f/I CD L
"~
0.6
Parameter: DMSO
0.4 0.2 0.0
con centration,
coNso 1% (v/v) rl o ~7 30 Zl 10 O 40 O 20 ' 0
9
I
I
i
l
2
I
i
.
3
I
i
4
I
5
i
I
6
9
i
7
.
8
Time, days
Figure 3" Deactivation of o~-chymotrypsin at 30 ~ and pH 7.5 The series-type of model did explain at least partly the observations with respect to the stability of ~-chymotrypsin. During thermal treatment a stable intermediate enzyme species (El) was formed the specific activity of which was higher than that of the initial species (E). The rate of formation, and particularly the rate of disappearance of this intermediate species, were influenced by the presence of DMSO. The rate of disappearance decreased at temperatures lower than 44 ~ in the presence of DMSO. At higher temperatures DMSO accelerated the rate of deactivation. Apparently, DMSO was able to stabilize o~-chymotrypsin by stabilizing the intermediate
227 species against further attack. The formation of the intermediate species was accelerated by increasing the temperature and the pH. At 50 ~ the reaction was so fast that the intermediate species could no longer be observed - as shown in Fig. 1. It is remarkable that there was only a small influence of pH on the stability of ~chymotrypsin in solutions containing 20, 30, and 40 % (v/v) DMSO at 30 ~ - as shown in Fig. 2 and 3. Table 1 summarizes the half-life of the initial enzyme species. These have been calculated in the case of experiments at 50 ~ as
In(2)
tl/2= k---~
(4)
In(2) tl/2 _ /" ,kl:~
andas
Table 1" Half-life of o~-chymotrypsin T
pH
,
for those at 30- 44 ~
(tl/2 / h) in the presence of DMSO
DMSO-concentration
/ % (v/v)
-
0
10
20
30
40
6.0 7.5 8.5 6.0 6.0 6.0
38.1 6.37 6.60 4.58 2.68 1.13
91.1 28.1 26.3 13.1 4.62 1.12 ,
181 114 110 35.3 6.42 0.88
242 185 268 79.1 n.d. 0.72
342 544 318 n.d. n.d. 0.35
~
30 30 30 39 44 50
(5)
.
,
The influence of DMSO on the main rate constant of deactivation (k2) is summarized in Fig. 4 for temperatures at which DMSO showed a stabilizing action. The activity ratio (~) was found to be normally about 1.3. Thus, the intermediate enzyme species seemed to be 30 % more active than the initial one. 10
-3 -
Parameter: pHi "7 r ...,.
E
--r'l--
- and
te mperature
8.5;
--Z~--7.5;
10-4
/ ~
30
--0--
6.0;
30
--~7--6.0;
30 39
e-
C
o
1 0. 5
1 0 .6
J
0
i
I
10
i
I
20
,
,
I
30
,
I ,,,
40
DMSO concentration, % (v/v)
Figure 4: Influence of DMSO on the main rate constant of deactivation
228
Finally, the activity of ~-chymotrypsin has been determined in the presence of DMSO and three different substrates in order to test, if a selective inhibition of proteolysis may eventually stabilize the enzyme against autohydrolysis. The data are gathered in Fig. 5. 120
lOO =~
~
80
~
60
~
4o
, O~ 0
rl
Parameter: s ubstrate - - / k - - SUPHEPA ~ - - 0 - AZOCASEIN - - E l - - BTEE
20 0
0
10 20 30 DMSO concentration, % (v/v)
40
Figure 5: Influence of DMSO on different reactions of ~-chymotrypsin Obviously, there was an inhibitory effect of DMSO on all of the reactions tested at least at high DMSO content. However, the effect seemed to be less pronounced and less specific for proteolysis - as tested by the solubilization of azocasein - in order to be able to interpret the stabilization of ~-chymotrypsin by DMSO as owing to the inhibition of autodigestion. Thus, the results have shown that soluble ~-chymotrypsin may be stabilized by DMSO. This was true even for alkaline conditions, a domain of particular interest for synthetic reactions. Even a continuous use of ~-chymotrypsin in ultrafiltrationmembrane reactors (Fiaschei eta/. 1982) may be envisaged. In addition, it would be of quite some interest to know, if this stabilizing principle may be found for other enzymes as well. REFERENCES Flaschel E., Wandrey C., Kula M.-R. (1982), Adv. Biochem. Eng./Biotech. 26, 73-142 Kisee H., Fujimoto K., Noritomi H. (1988), J. Biotechnol. 8, 279-290 Moresoli C, Flaschel E., Renken A. (1992a), Biocatalysis 5,213-231 Moresoli C., Zaza P., Flaschel E., Renken A. (1992b), Biocatalysis 5, 203-211 Pliura D.H., Jones J.B. (1980), Can. J. Chem. 58, 2633-2640
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
229
T h e r m a l stabilization by its ligands of NADP+-isocitrate d e h y d r o g e n a s e f r o m the thermophilic c y a n o b a c t e r i u m Phormidium laminosum* Miguel A. Pardo, Maria J. Llama and Juan L. Serra§ Enzyme and Cell Technology Group, Department of Biochemistry and Molecular Biology, University of the Basque Country, Faculty of Sciences, P.O. Box 644, E-48080 Bilbao, Spain
The thermostability of the NADP§ isocitrate dehydrogenase [Isocitrate: NADP § oxidoreductase (decarboxylating), EC 1.1.1.42] purified to homogeneity from the thermophilic non-N2-f'lxing cyanobacterium Phormidium laminosum was studied. The catalytic activity of the enzyme increased up to 55~ and the calculated activation energy (Ea) for catalysis was 9.5 +__0.1 kcal mol ~. In the absence of ligands the enzyme was fully stable until 50~ but more than 75% of its activity was lost when it was incubated for 15 min at 60~ and it was completely inactive at 650C. The thermal inactivation obeyed a first-order reaction and the calculated Ea for enzyme denaturation was 91.5 _ 0.1 kcal mol1. The presence of either D,Lisocitrate or NADP § did not stabilize the enzyme, but the combined presence of D,L-isocitrate and Mn 2§ increased significantly the enzyme thermostability and, at least, 50% of the activity remained after 15 min at 73~ The calculated Eo for thermal inactivation of the enzyme-D,Lisocitrate-Mn 2§ ternary complex was 109 +_0.1 kcal mol-~. These results together with the fact that the Km values of the enzyme for D,L-isocitrate and NADP § depended on the nature of the divalent metal cation used as a cofactor suggest that the D,L-Isocitrate-metal 2+ complex is the true substrate of the enzyme.
1. INTRODUCTION In cyanobacteria the NADP§ isocitrate dehydmgenase (NADP-IDH) is the enzyme responsible to provide the cell with 2-oxoglutarate which is the carbon skeleton acceptor of the ammonium resulting from nitrate assimilation. Since the tricarboxylic acids cycle is incomplete in these photosynthetic prokaryotes (due to the lack of both 2-oxoglutarate dehydrogenase and succinyl-CoA synthetase activities [ 1]) IDH activity plays a key role in the linking of nitrogen and carbon metabolisms [2]. Nitrogen starvation caused in the thermophilic filamentous non-N2-flxing cyanobacterium P. laminosum important changes in morphology and biochemical composition of cells accompanied with a noticeable increase on the cellular
* Supported by grants (DGICYT PB95-0509 and DGES PB96-1471) from the Spanish Ministry of Education and Culture. M.A.P. is the recipient of a scholarship from the Basque Government. To whom correspondence should be addressed
230 level of several enzymes involved directly in nitrate assimilation as well as in NADP-IDH activity. Although the cyanobacterial NADP-IDH has been characterized in Anabaena sp. PCC 7119 [3] and Anabaena cylindrica [4] and purified to homogeneity from nitrogen-sufficient cells of Anacystis nidulans [5], Synechocystis sp. PCC 6803 [6] and Anabaena sp. PCC 7120 [7] no data on the effect of temperature on the stability of the enzyme have been reported so far. In this paper we report the thermostability of the enzyme from the thermophilic cyanobacterium P. laminosum in the presence and absence of its ligands. The enzyme was purified to homogeneity from nitrogen-starved cells which showed 8 to 10-times higher specific activity than nitrate-grown cells. A purification step consisting in the incubation for 15 min at 65~ of the crude extract supplemented with D,L-isocitrate and Mn ~* appeared as a key step in the purification strategy.
2. MATERIALS AND METHODS 2.1. Materials Mono Q HR 5/5 column and Phenyl Sepharose CL-4B were from Pharmacia Biotech (Uppsala, Sweden). DEAE-ceUulose (DE-52) was from Whatman (Maidstone, U.K.). Reactive Red-120 Agarose and D,L-isocitrate were from Sigma Chemical Co. (St. Louis, U.S.A.). NADP was from Boehfinger (Mannheim, Germany). Inorganic salts and other chemicals were reagents of analytical grade supplied by E. Merck (Darmstadt, Germany).
Organismand cell culture Phormidium laminosum (Agardh) Gom. strain OH-I-p.Cll was originally obtained from Prof. R. W. Castenholz (University of Oregon, Eugene, OR, U.S.A.). Cells were grown photoautotrophicaUy [8] and maintained routinely in axenic monoalgal cultures at 45~ in liquid medium D supplemented with 0.5 g 1t NaHCO3 [9] which contained 10 mM nitrate as the sole nitrogen source. Cultures were stirred by an air stream and illuminated by white fluorescent lamps with a photon flux density at the surface of the vessels of about 100 I.tmol rn2 st. To obtain nitrogen-starved cultures, cells were grown as indicated above in glass containers with 60 1 of medium D containing only 1 mM KNO3 as nitrogen source [10]. Cells grew until nitrate exhaustion and then the culture was maintained in the nitrogen-free medium for further 60-65 h under the same light and temperature conditions used for growth. 2.2.
2.3. Enzyme assay NADP-IDH activity was routinely measured at 45~ by following the reduction of NADP + at 340 nm in 50 mM Tris-HC1 buffer, pH 7.0, containing 1.7 mM D,L-isocitrate, 0.2 mM NADP § and 2 mM Mn2§ 4 mM Mg 2+) in a final volume of 1 ml. The reaction was started by the addition of enzyme. One unit (U) of activity catalysed the appearance of 1 lamol NADPH2 per min under standard assay conditions. The affinity (Kin) of the enzyme for its ligands was determined in both the Mn 2§ and Mg2§ reactions. Kinetic parameters were calculated fitting data by non-linear regression to the Michaelis-Menten equation. Protein was determined according to Peterson [11] using crystalline bovine serum albumin as the standard.
231
2.4. Thermostability studies Thermal stability was determined by assaying at 45~ the residual activity of aliquots of the pure enzyme incubated for 15 min at different temperatures in the presence or absence of ligands. The activation energy (Ea) for the decarboxylating activity and for thermal inactivation of enzyme were calculated from the slope of Arrhenius plots of logarithm of catalytic or inactivation first-order rate constants versus the reciprocal of the absolute temperature. The slope corresponded to -Ed2.303 R, where R is 1.987 cal K t mol t.
2.5. Enzyme purification The enzyme was purified from nitrogen-starved cells of P. laminosum to electrophoretical homogeneity by an unpublished procedure which includes heat treatment and several conventional low-pressure chromatographic steps performed at 4~ followed by a final ionexchange step using FPLC at room temperature. Cells were harvested by centrifugation, washed and resuspended in 20 mM Tris-HCl buffer (pH 7.0) containing 0.5 mM EDTA and 8 mM 2-mercaptoethanol. Cells were disrupted by sonication (Soniprep 150, MSE, Mabor Royal, U.K.) in an ice-bath and the preparation was centrifuged at 12.000 x g for 60 min at 4~ Solid D,L-isocitrate (to 1.7 raM) and MnSO4 (to 4 mM) were added to the cell-free extract which was then incubated at 65~ for 15 min. The preparation was cooled and centrifuged at 11.000 x g for 60 min at 4~ The supematant was decanted and chromatographied through DEAE-cellulose (DE-52), Reactive Red-120 Agarose and Phenyl Sepharose CL-4B. Finally, a homogeneous IDH preparation was obtained after ion-exchange chromatography in a Mono Q HR 5/5 column.
2.6. Polyacrylamide gel electrophoresis (PAGE) analysis The progress of enzyme purification, the determination of molecular mass of the native enzyme and the formation of high-size aggregates of enzyme during its heat-denaturation in the presence of ligands were analysed by nondenaturing PAGE. Electrophoresis was run in a PhastSystem equipment using 8-25% (w/v) acrylamide gradient PhastGels (Amersham Pharmacia Biotech, Uppsala, Sweden) or 10% (w/v) homogeneous acrylamide gels in a BioRad (Hercules, U.S.A.) equipment. Protein bands were stained with 0.2% (w/v) Coomassie brilliant blue R-350.
3. RESULTS AND DISCUSION
3.1. Purification of NADP-IDH The purification protocol used in this work allowed the purification of enzyme from 600 to 650-fold to a specific activity of 500 U/mg protein with a final activity yield of 25%. This specific activity value is much higher than those reported for the enzyme purified from nitrategrown cells of Synechocystis (15.7 U mgt protein) [6] and Anabaena (25.0 U mg ~ protein) [7], and probably it represents the highest specific activity for a NADP-IDH reported so far for the enzyme isolated from photosynthetic organisms. The initial heat treatment of the crude extract at 65~ in the presence of D,L-isocitrate and Mn 2§ was crucial since it removed most of the chlorophyllous pigments present in the crude extract and increased by near 3-fold the NADP-IDH specific activity.
232
3.2. Affinity of NADP-IDH for its ligands The enzyme required for its full activity the obligatory presence of either Mg 2+ and Mn 2* as an essential activator, although other divalent cations were also partially effective. The K= values for D,L-isocitrate and NADP + calculated in the Mn 2. and Mg2+-dependent reactions were significantly different (data not shown). Moreover, the calculated K= for Mg 2§ (159 +_0.5 I.tM) was near 7-fold higher than that for Mn 2§ (24 • 3.0 I.tM) despite the fact that Mg 2§ is considered to be the NADP-IDH physiological cofactor [6]. 3.3. Effect of temperature on the activity and stability of NADP-IDH The effect of temperature on NADP-IDH activity was examined by monitoring the appearance of NADPH2 in the complete reaction mixture incubated at temperatures ranging from 25 to 65~ The activity increased linearly with temperature up to 55*(3. The Eo for catalysis calculated from an Arrhenius plot was 9.5 _ 0.1 kcal mol ~. The thermostability of the enzyme was studied in detail in the absence or presence of its ligands, all of them supplied at the same concentrations than those used in the standard assay for activity (Fig. 1). The enzyme showed similar stability against denaturation when incubated alone or in the presence of either NADP or D,L-isocitrate. The presence of Mg ~+ or Mn 2+ alone increased the enzyme thermostability, but in a lower extent than when each one of these cations was added in combination with D,L-isocitrate.
80-
40 "-i-Isocitrate
0
'
40
45
~~
~ ~
\ ! \
NADPornoligands ~ \ ,
,
50
55
\ ~
, -'~'~--f3-~O4
60
TEMPERATURE
65
70
~L 75
(~
Figure l. Effect of ligands on the thermostability of P. laminosum NADP-IDH activity. The pure enzyme in 50 mM Tris-HC1 buffer, pH 7.0, was incubated for 15 rain at the different temperatures in the presence or absence of ligands. After cooling the samples in an ice-bath the residual activity was assayed at 45~ using the standard assay. The concentrations used were: 1.7 mM D,L-isocitrate, 2 mM Mn 2+, 4 mM Mg 2. and 0.2 mM NADP. The effect of temperature on the denaturation rate of the enzyme was studied in the absence of ligands or in the presence of D,L-isocitrate plus Mn 2.. In the former case the P. laminosum NADP-IDH activity decayed following first-order kinetics (Fig. 2A). The Eo for thermal denaturation calculated from an Arrhenius plot (Fig. 2B) was 91.5 _.+0.1 kcal mol ~.
233
0.2
-0.6
---, 9 -0.2 O
-0.8
A
LU r 0
-0.6
--" -
o ._,1
-1.0
o0
-1.0
-1.2
-1.4 -1.8
B
,
0
4
.........
,
8
,
12
-1.4
16
2.96
I
I
I
2.98
3.00
3.02
TIME (min)
,,
3.04
1000FF (K -1)
Figure 2. (A) Denaturation time-course of P. laminosum NADP-IDH activity at various temperatures in the absence of ligands. The pure enzyme in 50 mM Tris-HC1 buffer, pH 7.0, was incubated for the indicated times at 58, 60 and 62~ Then, the residual activity of samples was assayed at 45~ in the standard reaction mixture. (B) Arrhenius plot of first-order denaturation constants (calculated from the slopes of Fig. 2A) against 1/T.
The presence of D,L-isocitrate and Mn2+ in the incubation mixture increased significantly the enzyme thermostability, and 50% of the initial activity remained after 15 min at 73~ The denaturation of the enzyme also obeyed first-order kinetics (Fig. 3A) and the E= for thermal inactivation of the enzyme-isocitrate-Mn ~-+complex was 109 _ 0.1 kcal mol = (Fig. 3B).
0.2 .......
~
~'0.-~ -0.2 UJ
r---i
iii
0.0 ~~0.~_.~~
B
7 2 '~2 A
-0.4
-0.6 -1.0
..J
o
-o.8 -1.2
-J -1.4 -1.8
!
0
4
I
!
8 12 TIME (rain)
16
-1.6 2.84
I
2.86 2.88 2.90 1000FF (K 4)
2.92
Figure 3. (A) Denaturation time-course of P. laminosum NADP-IDH activity at various temperatures in the presence of 1.7 mM D,L-isocitrate and 2 mM Mn2+. The pure enzyme in 50 mM Tris-HC1 buffer, pH 7.0, was incubated for the indicated times at 72, 74, 76 and 78~ Then, the residual activity of samples was assayed at 45"13 in the standard reaction mixture. (B) Arrhenius plot of first-order denaturation constants (calculated from the slopes of Fig. 3A) against lfr.
234 The size of aggregates resulting from thermal denaturation of enzyme in the presence or absence of ligands was analyzed by nondenaturing PAGE in homogeneous 10% (w/v) acrylamide gels. In all cases aggregates of very high molecular size appeared which were unable to penetrate into the separating gel (data not shown). Despite the fact that P. laminosum is regarded as a thermophilic cyanobacterium, the calculated values of Eo for thermal denaturation of enzyme either alone or forming complex with its substrates are lower than those reported for the NADP-IDH purified from the non-thermophilic fungus Cephalosporium acremonium [12] (E, for inactivation of 30.4 and 168.0 kcal mol "1 for the enzyme alone and with saturating concentrations of D,L-isocitrate and Mg 2+, respectively). However, the P. laminosum NADP-IDH activity was more thermostable than nitrate reductase (Eo = 33.5 kcal tool"1) [10] or nitrite reductase (Eo = 40.0 kcal mol1) [13] activities purified from this cyanobacterium. Taking into account that the affinity for D,L-isocitrate and NADP + of the P. laminosum NADP-IDH depended on the nature of the divalent cation used as an essential activator together with the fact that the ternary complex enzyme-D,L-isocitrate-Mn 2+ was more thermostable than the binary complexes of the enzyme with either D,L-isocitrate or NADP + did, the conclusion that the complex D,L-isocitrate-Metal 2§ is the true substrate of the enzyme can be drawn.
REFERENCES 1. R.Y. Stanier and G. Cohen-Bazire, Ann. Rev. Microbiol., 31 (1977) 225. 2. J. Pearce, C.K. Leach and P. Gadal, J. Gen. Microbiol., 55 (1969) 371. 3. L. Kami and E. Tel-Or, In: Photosynthetic Prokaryotes (G.C. Papageorgiou and L. Packer, eds.) pp. 257, Elsevier Science Publishing Co, New York, 1982 4. H. Papen, G. Neuer, M. Refaian and H. Bothe, Arch. Microbiol., 134 (1983) 73. 5. G.M. Friga and G.L. Farkas, Arch. Microbiol., 129 (1981) 331. 6. M.I. Muro-Pastor and F.J. Florencio, Eur. J. Biochem., 203 (1992) 99. 7. M.I. Muro-Pastor and F.J. Florencio, J. Bacteriol., 176 (1994) 2718. 8. M.I. Tapia, M.J. Llama and J.L. Serra, Planta, 198 (1996) 24. 9. R.W. Castenholz, Schweiz Z. Hidrol., 32 (1970) 538. 10. F. Blanco and J.L. Serra, In: Metabolismo del Nitr6geno (M. Pineda and F. Castillo, eds.) pp. 68, SEB, C6rdoba, 1992 11. G.L. Peterson, Methods Enzymol., 91 (1983) 95. 12. J. Olano, D. de Arriaga, F. Busto and J. Soler, Appl. Environ. Microbiol., 61 (1995) 2326. 13. J.M. Afizmendi and J.L. Serra, Biochim. Biophys. Acta, 1040 (1990) 237.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
235
The characterization of protein stability from hydrogen exchange 9 Maximum entropy analysis of co-solvent effects on exchange rates Badredine Souhail, Isabel M. Plaza del Pino and Jose M. Sanchez-Ruiz Departmento de Quimica Fisica (Facultad de Ciencias) e Instituto de Biotecnologia. Universidad de Granada. 18071 Granada, SPAIN. The overall stability of a protein, as well as the stability of its specific regions, may, in principle, be characterized from the analysis of cosolvent (denaturant) effects of hydrogen exchange rates (1). This type of analysis presupposes however, that the protein can only'exist in a given number discreet states, an assumption which, according to the so called new-view of protein folding, might not hold. We show here that it is possible to pose the analysis of the cosolvent dependence of the hydrogen exchange rates in terms of continuous distribution of rates. In addition, we also demonstrate that this distribution may be determined from the experimental data by using a Maximum Entropy algorithm. Finally, we describe the application of this approach to two series of hydrogen exchange data corresponding to cytochrome c (1) and ribonuclease H (2). THE MAXIMUM ENTROPY METHOD Let P(t) be a certain property of protein (measured experimentally or calculated from experimental data).We assume that the property decreases monotonically with t, and that its variation can be described in terms of continuous distribution: +oo
PC"t(t)=fj(ln~.)
e-x, d0n;t)
-oo
wheref(ln2) is the amplitude distribution function and ~, is the rate constant. pca~(t) is the calculated property, and the calculated distribution must: l) Fit the experimental data according to the Least Squares Criterion. 2)Contain a minimum information according to the Maximum Entropy Criterion. The most probable distribution, is a compromise between the two criteria, and is obtained by maximizing : ~2 _ o~S
236 where 2'2 is given by the Least Squares Criterion:
X2 :
i=N ( p e x p ~ [ --i =0
__ p cat
--i 0i
J
S is the Shanon Entropy of the distribution:
s = - f f tnf dt~f
and the value of ct is found from Bayesian Statistics arguments (3). In order to apply the Maximum Entropy Method (MEM) to the analysis of hydrogen exchange in proteins, some transformations of the exchange equations are necessary. Exchangeable amide hydrogens (NH) that are involved in hydrogen bonded structure can exchange with solvent hydrogens only when they are transiently exposed to solvent. In most cases amide hydrogen exchange in proteins conforms to the EX2 mechanism (4), and the exchange rates are analyzed according to the following scheme: Rop
closed
~,
Rex
open
---
exchange
The equilibrium opening constant corresponding to one mechanism (i) for one hydrogen is given by the equation: AGi gop
,i
= e
RT
If the exchange occurs by various mechanisms, then the effective opening constant is given by the equation: i =n
Kop,eff = ~ e
i=0
AG~ Rr
237 The dependence of the unfolding free energy on denaturant concentration is assumed to be linear: A G , : AG,W-m[C]
: AGT+m([C*]-[C] ) = AG,*+mAC
AGT:AG, w- m[C*]
With
; AC:[C*] - [C]
where A G 7 is the Gibbs energy change extrapolated to zero concentration of denaturant, C is the concentration of denaturant, and C* is a given concentration taken as reference. The slope m depends on the additional denaturant sensitive surface exposed in the unfolding mechanism. The opening reactions that determine protein hydrogen exchange may expose much or little new surface. Then, the effective opening constant can be expressed by: AGi" -
K o p eft = ~ , t
e
AC
AC
- m - -
Rr e
-m
Rr : ~ K~p., e
Rr
i
We now take the above equation ~o the continuous limit, oo
f,
Kopxff = gop(m ) e
_
mAC
-
Rr d m =
0
mAC
f,
Kop(ln(m)) e
Rr dOn(m))
-~
where Kop* is the amplitude distribution for the effective opening constant of the hydrogen exchange. Now we can apply the MEM approach by using the following identification: Kop.eff= P
9 '
~ = m
;
AC t
-
RT
We show in figure 1 the applicability of the Maximum Entropy approach to the hydrogen exchange in cytochrome c and ribonuclease H. Thus, figure 1A shows the variation of e -Ac/Rr versus A C / R T corresponding to Isoleucine-25 and Isoleucine-78 of the Ribonuclease H (2), and figure 1B shows the calculated distribution of K,,p versus ln(m), derived from maximum entropy analysis. The distributions show one peak in the case of Isoleucine-25, and two peaks for the Isoleucine-78. The number of peaks in the
238
distribution corresponds to the number Of a different opening reactions that allow hydrogen exchange to occur.
4000
A
e,
2000 ll
1 2 AC/RT (M kJ "1 mol "l)
1200
t
il
E
i'
B
800 -
iI
'1
!
|1 I I
'I o
|l
400 "
|| I I
't
| |
-4
-2
0 Ln(m)
2
4
F i g u r e 1. (A) Dependence of opening constant with the concentration of Guanidine. (e) Isoleucine-78 and (I)Isoleucine-25, the continuous lines(---) are the fitting of the experimental data (data are taken from Ref. 2), derived from Maximum Entropy analysis. (B) Examples of the distributions calculated from the maximum entropy analysis of the opening constant. The continuous line corresponds to Isoleucine-78, and the dashed line to Isoleucine-25.
239 Finally in order to ascertain the statistical errors involved in this Maximum Entropy analysis, 20 Monte Carlo simulations of the data were generated and subjected to Maximum Entropy analysis. Examples of the resulting distributions are shown in Figure 2. Peak positions and peak areas are given in Figure 3.
500 400 -g
A
300 200 100 0
! 1
i
! 2
i
i
iii
!
!
3
4
i
! 5
in ( k J m o l "1 M -1) 300 -
B 200 .g
100
0 0
1
2
3
4
5
6
m ( k J m o l q M "l) F i g u r e 2. Examples of the distributions derived from Monte Carlo simulations of the hydrogen exchange data. Isoleucine-25 (Fig. 2A) and Isoleucine-78 (Fig. 2B).
240
i
5-
4 E E
A
m
m
lle|.."
3
2
#
m
,
mOe.lllllle ,
m
_-_- ;_---_
=,,,= = ;
m
1 -
0
n l
m
-
= ~--
l
ml
0
mm
5
_- ; -
10
mm
15
NUMBER OF MONTECARLO olml
m
20
SIMULATION
200 B
150e=
"-
< i.=
<
100m
50- I 0
m I
1
I
i
I
I
2 3 4 m (kJ m o l "l M "l)
I
5
6
Figure 3. (A) Positions of the peaks observed in the distributions calculated from the Maximum Entropy analysis of 20 Monte Carlo simulations of the hydrogen exchange data. (B) Plot of peak area versus peak position. Peak area were calculated from the numerical integration of the distributions.(o ) Isoleucine-25 and (m) Isoleucine-78. ACKNOWLEDGMENT This work was supported by grant from the Fundaci6n Ram6n Areces. REFERENCES 1. Y. Bai, T. R. Sosnick, L. Mayne, S. W. Englander, Science, 269 (1995) 192-197. 2. A.K. Chamberlain, T. M. Handel, S. Marqusee, Nat. Struct. Biol., 3 (1996) 782-787. 3. J. Skilling. in Maximum Entropy and Bayesian Methods (1989) (Skilling, J., Ed.), pp. 45-52, Kluwer, Dordrecht, The netherlands. 4. S.W. Englander, N. R. Kallenbach, Q. Rev. Biophys. 16 (1983) 521-655.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Preferential h y d r a t i o n changes mixtures.
upon protein unfolding
241
in water-cosolvent
Hassan O. Hammou, Isabel M. Plaza del Pino and Jose M. Sanchez-Ruiz ~ ~Facultad de Ciencias, Departamento de Quimica Fisica 18071-Granada, Spain. E-mail: [email protected] We have characterized the unfolding energetics of ribonuclease A (RNase) and hen-eggwhite lysozyme as a function of temperature, pH and concentration of several cosolvents (sucrose, glucose, glycerol, polyethylenglycol 8000) (Table I, Table II) that are expected to be preferentially excluded from the surfaces of proteins, and we have calculated the corresponding unfolding changes in preferential hydration (AF2,)(Table III) [Timasheff (1993) Annu. Rev. Biophys. Biomol. Struct. 22, 67-97; Plaza del Pino and Sanchez-Ruiz (1995) Biochemistry 34, 8621-8630]. We find that these unfolding changes in preferential hydration are significantly smaller than several plausible estimates of the unfolding change in the number of water molecules required for first monolayer coverage of the protein (Table IV), even when, for the purpose of this calculation, the solvent accesibility of the unfolded state is modeled on the basis of compact fragments excised from native protien structures [Creamer, Srinivasan and Rose (1997) Biochemistry 36, 2832-2835]. An analysis in terms of the two-domain (local -bulk) solvent model [Record and Anderson (1995) Biophys. J. 68, 786-794] shows that the low values found for unfolding changes in preferential hydration could be the result of the entrance of rather small amounts of cosolvent in the local domain (the protein domain, that is, the solvent "shell") of the native and/or the unfolded protein. The two-domain model suggests that small differences in the quantities of cosolvent present in the local domain can determine if a certain substance is a proteins stabilizer or an agent denaturant (Figure 1). In general, the two-domain model suggests that even a weak protein-cosolvent interaction may significantly distort the membrane-free, osmotic stress estimates of the number of the water molecules involved in protein conformational change.
1. A TWO-DOMAIN SOLVENT MODEL... Two solvent domains are assumed: 1) The bulk domain. 2) The protein domain, that is, the solvent "shell" affected by the protein. The number of molecules of water (1) and cosolvent (3) present in the protein domain (i,e., the number of solvent molecules "bound" to the protein) are termed B, and B3. It can be shown that the denaturational change in preferential hydration is related to the denaturational changes in actual water and cosolvent binding by:
242
AF2t = ABt -
10: ) M~ m 3 - A B3
(1)
where M! is the molecular weight of water and m3 is the cosolvent molality.
2. PLUS A SOLVENT-EXCHANGE MODEL When a eosolvent molecule enters the solvent shell surrounding the protein, v molecules of water are released to the bulk solvent domain. Then: AB1 = AB ~ - v_AB3
(2)
where AB~ is the denaturational change in actual water binding that would be observed if no cosolvent binding took place. AB~ is the higher possible value for the denaturational change in preferential hydration and, on the basis of aecesible surface area values, has been estimated to be about 400 molecules of water per mole of protein. Upon substitution of eq 2 into eq 1, we obtain:
AF21 = AB ~ - v +
Ml_m3
AB3
(3)
For m3=l molal, 103/M~-m3 is 55.55, which is expected to be significantly higher than the exchange stoiehiometry (v). Table I Comparison between the energetic unfolding parameters derived from the analysis of DSC thermograms and optical unfolding profiles. DSC Protein
Cosolvcnt
pH
Tm (~
optical r~
(kJ/mol)
H~ (kJ/mol)
Tm (~
H~H (kJ/mol)
H vH
RNase
none
4.02
56.4
349
355
1.02
55.8
347
RNase
sucroseI M
3.60
61.3
394
370
0.94
60.6
387
RNase
glucose2M
3.54
64.2
419
380
0.91
63.9
398
RNase
glycerol 4M
3.55
59.4
371
399
1.08
59.2
361
RNase
PEG 25%
3.55
54.9
319
340
1.07
54.4
337
lysozyme
none
3.01
72.5
500
496
0.99
71.1
469
lysozyme
glucose2M
2.20
72.0
520
509
0.98
71.8
518
lysozyme
glycerol 4M
2.18
65.11
527
452
0.86
64.2
456
490
1.08
54.9
403
lysozyme PEG 25% 2.30 54.8 452 ~ Calorimetric to van't Hoff enthalpy ratio: r=AH~/AI-FH.
243 Table II Unfolding heat capacity changes and unfolding enthalpies at the reference temperature (50~ for RNase 65~ C for lysozyme) derived from global analyses of optical unfolding profiles, a
PROTEIN
RNase
lysozyme
COSOLVENT
ACp (kJ/K.mol)
AHR (kJ/mol)
none sucrose (0.25-1 M)
4.004-0.70 (4.7) 4.104-0.45
339+3 3474-6
glucose (0.5-2 M)
4.86+0.57 (3.2)
341+4
glycerol (1-4 M)
4.75+0.40 (4.6)
351+6
PEG 8000 (10-26%)
3.864-0.86(4.2)
350~3
none
6.38+ 1.10 (5.6)
439+ 11
glucose (0.5-2 M)
6.01+0.83 (5.1)
463+8
glycerol (1-4 M)
5.23+ 1.16 (5.9)
4794-16
PEG 8000 (10-25%)
7.05+0.97 (5.3)
478+19
a Each value is the average of the results obtained from the global analysis of several (usually four) different data sets of six or seven unfolding profiles each. No significant cosolvent concentration effects were detected. Therefore, for each given cosolvent results corresponding to different cosolvent concentrations were included in the calculation of the average value. The unfolding heat capacity changes given in brackets were derived from the analysis of DSC thermograms carried out at the highest cosolvent concentrations employed (1 M sucrose, 2 M glucose, 4 M glycerol, 25% PEGsooo).
244 Table HI Unfolding changes in protein-cosolvent preferential interaction parameter (kJ.kg-mol 2) and preferential hydration (mol-water/mol-protein) at 25 ~ PROTEIN RNase
lysozyme
COSOLVENT
A(~23
~k~"21
A(~23
A~"2!
sucrose 0.6 mol/kg
8.04•
157•
glucose 1.3 mol/kg
4.41•
92•
4.06•
85•
glycerol 2.8 mol/kg
1.43+0.06
30• 1
1.51 +0.32
32+7
PEGs0oo 15 %
-0.05+0.09
-14+26
-0.33•
97•
' The uncertainties given are derived from the linear fit employed in the temperature extrapolation of A~23to 25 ~ The actual error associated to the values given in this Table may be larger.
Table IV Theoretical estimates of the unfolding changes in accesible surface area (A2) and the number of water molecules in contact with the protein surface (mol-water/mol-protein). ~ PROTEIN RNase
lysozyme
MODEL FOR THE EXPOSURE TO SOLVENT IN THE UNFOLDED STATE
AASA
Z~N,
AASA
AN,
tripeptides
14256
15437
extended conformation (upper bound)
10498
634 (1782) 467 (1312)
686 (1930) 501 (1408)
fragments from folded structures (lower bound)
6422
285 (803)
7229
11262
321 (904)
a AASA values are calculated as ASAU-ASAN, where ASANvalues were taken from Miller, Janin,
245 Lesk and Chothia, (1987) J. Mol. Biol. 196, 641-656. and the ASA v values for the several models [Creamer, Srinivasan an Rose, (1995) Biochemistry 34, 16245-16250, Creamer, Srinivasan an Rose, (1995) Biochemistry 36, 2832-2835] were calculated from the sequence by using programs made available by Rose and cols. 0attp://cherubino.med.jhmi.edu/~-folded/). AN~ values are calculated by using: AASA AN1 = ASAw
(4)
with ASAw equal to 22.5 A 2 and 8 A s (values in brackets).
15
urea
O
[-, <
10 glycerol
II0%
.......
~o
o
glu se
c~
I
sucrose
.,..
-
--., i
<1
N
| ; cD cD
"6 @ ,.o
O "i::
0
Figure 1" Theoretical estimates of unfolding change in the number of molecules of cosolvent in the local domain for the proteins indicated and in water-cosolvent mixtures of 1 molal concentration of cosolvent.
ACKNOWLEDGMENT This work was made possible by the support of the Fundaci6n Ram6n Areces.
a This Page Intentionally Left Blank
Stabilityand Stabilization of Biocatalysts
247
A. BaUesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Characterizing protein-cosoivent interactions coupled to protein refolding by kinetic calorimetry. Mohammed C. Boulaich, Antonio Parody-Morreale and Jos6 M. Sb.nchez-Ruiz.
Departamento de Quimica Fisica (Facultad de Ciencias) e Instituto de Biotecnologia. Universidad de Granada. 18071-Granada. SPAIN
The kinetics and thermodynamics of the refolding of hen egg-white lysozyme in concentrated guanidine solutions, have been characterized by using a titration microcalorimeter to monitor directly the rate of heat production or absorption as a function of time (Figure 1) (Figure 2).
1,9e-6
1,2e-6
5,0e-7
-2,0e-7 0
500
1000
1500
2000
T i m e (s)
FIGURE 1: The dashed line represents a typical calorimetric trace (heat flow versus time) obtained after addition of unfolded lysozyme (10/.tL, 3.3 M guanidine, pH 2. O) to the calorimetric cell (203 l.tL) containing guanidine (3.3M, pH 4.5). The total area under the curve divided by the amount of the folded protein in the cell is equal to the change (AH) for the reaction, while the time decay of the rate of heat flow depends on the kinetics of the reaction. This experiment was performed in 10 mM sodium acetate, pH 4.42, 3.3 M guanidinium hydrochloride at 25 ~ The solid line represents the same calorimetric trace after correction for the instrumental time response.
248
1,8e-6
1,3e-6
8,0e-7
3,0e-7 -
-2,0e-7 .... 0
500
I
I
1000
1500
....
2000
Time (s) FIGURE 2: Time dependence of the heat flow evolution observed upon pH-change to induce the protein refolding reaction (final pH, 4. 42) in the solutions of Lysozyme (20 mg/mL in Acetate buffer, 3.3 M guanidine, pH 2.0) by addition of Guanidine (3.3 M in Acetate buffer, pH 4.5). The rate constants (k) and the enthalpy change (zlt-I) were determined by non linear least-squares analysis of the equation (1) to the experiments data dQ _ rid-1 V k [Co] exp(-kt) dt
(1)
where [Ca] is the initial concentration of the unfolded protein, and the V & the volume of the calorimetric cell.
Refolding was initiated by a pH jump (from 4.5 to 2.0) and experiments were carried out at several temperatures and guanidine concentration.This technique allows us to determine both the energetics ( enthalpy and heat capacity) (Figure 3) and the kinetics (rate constants) of the refolding process at high denaturant concentration, data which are very difficult to obtain by using conventional calorimetric approaches (Privalov et al., J.Mol.Biol., 1992, 491-505).
249
180 0 140 -
0
t
II o
100
-
0 1"!
<1
!-1 60-
20 3,0
I
I
I
3,2
3,4
3,6
,,
3,8
[GdnHCI] (M) FIGURE 3: Guanidine concentration dependence of the enthalpy change for lysozyme refolding (10 mM sodium acetate, pH 4.3) at several temperatures: (s 18 ~ (0) 25 ~ (0) 30 ~ Error bars represent the associated standard errors, as given by the MLAB program using the diagonal elements of the covariance matrix.
The validity of this approach is supported by the excellent agreement found between the folding-unfolding rate constants obtained from the kinetics-calorimetry experiments and those derived from fluorescence measurements (Figure 4) (Ibarra-Molero and Sanchez-Ruiz, Biochemistry 1996, 14689-14702).
250
-3
o o
O
-5
#
25~ ~-7 "7
I
v
,
O
I
O
.I
I
t--"
%
"-3
9 O
O O
-5 30~ -7 I
0
2
.
O I
I
I
4
6
8
[GdnHCl] (M)
FIGURE 4: Chevron plots of folding-unfolding rate constant for lysozyme versus guanidine concentration for several temperatures. Empty symbol corresponds to kinetic experiment performed in the folding direction using the calorimetry. The data represented by black filled circle are takenfrom the equilibriumfluorescence intensity study at 30 ~ of lbarra-Molero and Sanchez-Ruiz (Biochemistry, 1996, 14689-14702).
ACKNOWLEDGMENT This work was made possible by the support of the Fundaci6n Ram6n Areces. REFERENCES 1. Ibarra-Molero, B., & Sanchez-Ruiz, J.M. (1996) Biochemistry 35, 14689-14702. 2. Makhatadze, G.I., & Privalov, P.L. (1992)J.Mol.Biol.226, 491-505.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
251
Characterizing cosolvent effects on protein stability: cold-denaturation of ubiquitin in the presence of guanidine Beatriz Ibarra Molero and Jose M. Sanchez-Ruiz Departamento de Quimica-Fisica (Facultad de Ciencias), Instituto de Biotecnologia Avd. Fuentenueva sn, 18071-Granada, Spain
We have recently characterized the thermal unfolding of ubiquitin from bovine red blood cells within the range 0-3.82M guanidine by Differential Scanning Calorimetry (DSC) experiments. At concentrations above 3M the high-temperature tail of the cold denaturation transition is observed indicating that ubiquitin also denatures upon cooling under these conditions. Fits to the data according to a two-state model were always excellent as shown in Figures 1 and 2.
10 kJ-K" 1.mol- 1
20
40 60 80 100 Temperature (~
120
Figure 1. DSC profiles for ubiquitin unfolding in 10 mM acetate buffer, pH 4.0 at different concentrations of GdmC1 shown alongside the profiles. The solid lines represent the best fit to a two-state model of the experimental data shown in open symbols. Curves have been shifted along the y-axes for display purposes.
252
_~ I
-40
~ v~ l ~ l a3.82M t -1 -1 I
I
.... I
0 40 80 T e m p e r a t u r e (~
.... 120
Figure 2. Temperature dependence of the heat capacity of ubiquitin in solutions containing different concentrations of GdmCl, showing both the heat and cold denaturation transitions. The solid lines are the best fit to a two state model of the experimental data shown in open symbols. The complete peak due to cold denaturation is shown for illustrative purpose only.
We compare our AH data at a given temperature and guanidine concentration with the temperature dependence of AH for ubiquitin reported by Wintrode et al. [ 1] (see Figure 3). These authors carried out DSC experiments in aqueous buffers at different pHs within the range 2.0-4.0. Assuming no pH effect, they showed the enthalpy change as a nonlinear function of the temperature, being these values higher than those in the presence of guanidine. This fact could be explained by the preferential binding of guanidine, to the denatured state, and the negative enthalpy associated to this process. We also show in Figure 3 the enthalpies of ubiquitin unfolding calculated according to the denaturant binding model [2], in which it is assumed that the GdmC1 molecules bind to both native and unfolded states of a protein presumably through hydrogen bonding. Using this model AHGa~c, (T) can be presented in terms of the enthalpy changes that result from GdmC1 binding to ubiquitin upon unfolding as follows :
253
A H~,~c, (T) = A n~,~c!
a h~c, -
Ka,~c, _ [GdmCl] - 1 + K~a~cl _ [GdmC1]
(1)
where An~amc~is the difference in the number of GdmC1 binding sites between unfolded and native states, Ahr,~a is the enthalpy of GdmCl binding to the protein, and K a~c~ is the GdmC1binding constant. In this equation Anr~c~ was taken to be equal to -15 [4], and the values for Ahramc~(25~ and K6,~,,c~(25~ were -11 J.mol ~ and 0.6 M ~, respectively [2]. Based on the previous estimates [4], the temperature dependencies of these parameters were taken to be 0.075 J.K~.mol ~ for Ah~amc~and 0.0075 K~.M ~ for Kr,~c 1. The adequacy of the denaturant binding model was tested by comparing the experimentally observed enthalpy of ubiquitin unfolding in the presence of GdmC1 with the predicted enthalpy, AHBM(T). The latter value was calculated from the ubiquitin unfolding data [1] in the absence of GdmC1, AH0(T), and the enthalpy of GdmC1 binding, AHG~cl(T ), as:
(2)
A H BM (T) = A Ho + A HG,~c, (T)
The agreement between the experimental and calculated results is remarkable indicating the adequacy of the binding model, at least as a phenomenological description of the guanidine effect on protein stability.
300
- o-
200
100
20
40
60 Temperature
80
100
(~
Figure 3. Temperature dependence of the enthalpy of ubiquitin unfolding. Solid squaresenthalpies of ubiquitin unfolding obtained from the fit of the curves presented on Figures 1 & 2; open circles-enthalpies of ubiquitin unfolding calculated according to equation (2); open squaresenthalpies of ubiquitin unfolding in the absence of GdmC1, taken from [ 1]. Solid lines represent polynomial fitting of the data points.
254
An interesting result is obtained if we compared the ACp data reported by Wintrode et al. with ours (see Figure 4). Thus, the ACp values reported by these authors were calculated as: ACp = C~ - C~
(3)
where Cp,N is the heat capacity of the native state (linearly extrapolated from low temperature data when necessary) and Cp,U is the heat capacity of the unfolded state computed from the aminoacid composition as [3]: N
C~ = (N- 1) _ Cp (-CHCONH) + ~
Cv (- Ri)
(4)
i=!
Clearly, the unfolding heat capacity changes calculated by using equations (3) and (4) should be interpretated as the values corresponding the "complete" unfolding of the protein at zero guanidine concentration.
OM 3.50M I
0
4
~.,.q
3.00M
2
0M
2.04M 1.01M
0
20
40 60 80 t00 Temperature (~
120
Figure 4. Temperature effect on ACp values. The hollow symbols are taken from the DSC study of Wintrode et al. [ 1] at different pHs in aqueous buffer conditions. The solid line represents the fit of a quadratic polynomial to the data. The filled symbols represent the data derived from our DSC experiments in the presence of different guanidine concentrations (shown in figure). Dashed line is the linear fit to our data.
255 As it is shown in Figure 4 at high temperatures there is an excellent agreement among the two series of data indicating that no guanidine effect is taking place and ACp is changing only due to a temperature effect, at least in this concentration range. However, a significant difference is found at high guanidine concentration. In fact, we obtained lower ACp values at high guanidine concentration. This is a surprising result since evidence for the opposite behavior has been reported by Makhatadze and Privalov [2]. The simplest explanation for the discrepancy would be that, at high guanidine concentration and room temperature, the amino acid residues are not fully exposed to the solvent in the unfolded state, perhaps reflecting a restriction in the conformational freedom of the polypeptide chain caused by the "binding" of the denaturant molecules. Of course, other explanations may also be aduced; for instance, the lower heat capacity change in the presence of guanidine could result from a guanidine-induced alteration in the thermodynamic properties of water that determine the hydrophobic interaction. Also, the possibility that the lower ACp value be an artifact caused by deviations from the two-state mechanism in the cold denaturation cannot be completely ruled out, since only the hightemperature tail of the cold denaturation transition is apparent in our thermograms (Figure 2). Clearly, the discrepancy shown in Figure 4 requieres further investigation.
ACKNOWLEDGEMENT This work was supported by Grant PB96-1439 (DGES, Spanish Ministry of Education and Culture) and the Fundacirn Ramrn Areces.
REFERENCES
1. Wintrode, P. L., Makhatadze, G. I., Privalov, P. L. (1994) Proteins: Struct., Funct., Genet. 18, 246-253. 2. Makhatadze, G. I. and Privalov, P. L. (1992) J. Mol. Biol. 226, 491-505. 3. Makhatadze, G. I. and Privalov, P. L. (1980) J. Mol. Biol. 213,375-384. 4. Makhatadze, G. I., Lopez, M., Richardson, J. M. and Thomas, S. K. (1998) Protein Science 7, 689-697.
a This Page Intentionally Left Blank
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Acetylcholinesterase-ethanol ligand exclusion
interactions: inactivation, substrate
257
and
F. Ortega, D. Garcia and J.-L. Marty Centre de Phytopharmacie, UA CNRS 461, Universit~ de Perpignan, 52, avenue de Villeneuve, 66860 Perpignan, France. 1. I N T R O D U C T I O N I n t e n s i v e use of o r g a n o p h o s p h a t e (OP) insecticides h a s led to m a n y ecotoxicological problems. Their p r i m a r y toxicity arises from the irreversible inhibition of a nervous system essential enzyme, the acetylcholinesterase (ACHE, EC 3.1.1.7). Phosphorylation of the active-site serine blocks the hydrolysis of the n e u r o t r a n s m i t t e r acetylcholine [1] and hence, causes the insect's death. However since cholinergic transmission is well conserved, these compounds are potentially toxic for all animals. To m o n i t o r the levels of insecticides in w a t e r and soil s a m p l e s , an a d v a n t a g e o u s a l t e r n a t i v e for rapid and sensitive detection of OP is t h e d e t e r m i n a t i o n of these insecticides using ACHE. The main drawback of such biological approach is the use of solvents for the extraction and concentration of OP, which can alter the enzyme properties [2]. In order to facilitate a b e t t e r insight into the solvent-induced modifications of ACHE, the purpose of this work was to determine the effect of a water-soluble solvent, ethanol, on ACHE. The question has been tackled in two major parts: 1) the effect of ethanol on the functional stability of AChE and t h a t of additives to stabilize the enzyme, and 2) the influence of ethanol on the AChE-catalyzed hydrolysis of the acetylthiocholine (ATC) s u b s t r a t e and on the inhibition of ACHE* by the OP paraoxon (PX) in addition to t h a t by the competitive reversible cationic inhibitor, e d r o p h o n i u m (Edro).
* AChE from Drosophila melanogaster was kindly provided by Pr D. Fournier (Universit~ de Toulouse, France).
258
2. INACTIVATION AND STABILIZATION OF AChE IN THE P R E S E N C E OF E T H A N O L The effect of ethanol on the stability of the AChE has been investigated as a function of ethanol concentration. Upon exposure to significant amounts of the water-miscible organic solvent, the enzyme inactivated in a multistate manner prior to the rapid and total loss of enzyme activity at ethanol contents above 20% (v/v) (figure 1). Our results were analyzed according to a serial-inactivation mechanism of the t y p e : E - ~ E, ~ E~ where E represents the initial enzyme activity at time zero, E1 is a partially active form an E2 is a totally inactive conformation [3].
T none, 15%(v/v) EtOH
~075
-w-4 ~
9 none, 20%(v/v) EtOH 0,5
Edro 200 ~M, 15%(v/v) EtOH
,v,,~
o Edro i mM, 20%(v/v) EtOH
0,25
30 60 90 120 150 180 Preincubation time (min) Figure 1: Kinetic of AChE inactivation vs ethanol concentration and edrophonium stabilization, at pH 8.0, 30~ The inactivation was started by the addition of AChE solution (50 nM). At regular time intervals, aliquots were withdrawn from the incubation mixture and the enzyme activity was measured. Activities have been normalized as fraction of the activity measured at time zero. The kinetic parameters determined with this model clearly indicated a decrease in the residual activity of the intermediate E1 form together with a concomitant increase in the inactivation rate of the first step upon gradual addition of the cosolvent to the medium. In an additional experiment, we showed that the presence of a reversible cationic inhibitor, edrophonium that is selective for the enzyme active-center antagonized both the initial unfolding of the native enzyme and the inactivation of the E1 transient (figure 1). Similar protection of AChE is provided by others cationic active-site related ligands such as propidium and tetrabutylammonium. For comparison, non-specific ligands (e.g. NaC1, polyethylene glycol, glycerol) afforded none or poor protection to AChE (data not shown).
259 3. E F F E C T O F E T H A N O L ON A C h E R E A C T I O N AND I N H I B I T I O N We investigated the influence of ethanol on both the AChE-catalyzed hydrolysis of ATC and on the inhibition of AChE by PX. E+AX( Kd > E . A X k2 > E A + X k3 > E + A (1) In scheme 1, AX is ATC or PX, EA the acetylated or the phophorylated ACHE, Kd the dissociation c o n s t a n t AChE-ATC or AChE-PX (M), k2 the acylation or phosphorylation r a t e c o n s t a n t s (min-1) and k3 t h e d e a c y l a t i o n or d e p h o s p h o r y l a t i o n r a t e c o n s t a n t (min-1). In the case of PX, k3 = 0 and the bimolecular inhibition constant ki (M-lmin -1) is defined as ki=k2/Kd.
The hydrolysis of ATC by AChE was monitored in the presence of different a m o u n t s of ethanol. The Lineweaver-Burk plots obtained from our results are shown in figure 2. 15
EtOH
Km
% (v/v) !
=L 9~ +
r--
II
0.0
40.7
9
5.0
41.4
9
10.0
53.3
9
12.5
60.8
[-q
15.0
111.8
0
17.5
121.1
/~
20.0
170.7
10
5
>
--
0
0
20
40 I/[ATC] (mM-l)
60
80
Figure 2" Lineweaver-Burk plots of AChE-catalyzed hydrolysis of ATC at various ethanol concentrations, pH 8.0, 30~ It can be seen t h a t 1) the double-reciprocal plots are linear all over the range tested for ethanol i.e. up to 20% (v/v) and 2) the lines cross to a common intercept indicating that the apparent maximal rate of the reaction is not affected by the organic solvent. Therefore, ethanol mainly appears to modify the affinity of AChE for ATC, as shown by the calculated values of Km (cf. figure 2). This a p p a r e n t competition between ethanol and ATC during the reaction of AChE has been described previously by Ronzani [4] both for ethanol and for several watermiscible solvents. However, Km is a composite constant t h a t includes both binding and kinetic components. Ethanol can undergo nucleophilic competition with w a t e r for deacylation of the acyl-AChE intermediate (equation 1), thereby possibly leading to a change in the rate limiting step of the catalytic process and hence, to a change in the significance of Km:
260
K
m
k 3 Kd = ~ =
k2+k3
k 3 Kd ~
k2
or
-~ K d
when
k2 > k3
or
k3< k2, respectively.
(2)
Therefore, the analysis of the system with ATC is far from straightforward and requires further arguments to comprehend the effect of ethanol on the enzyme reaction. Towards this goal, we investigated the influence of the solvent on the competitive inhibition by edrophonium of the enzymatic hydrolysis of ATC. As shown in figure 3, the value of the dissociation constant for the AChE-Edro complex (Ki) dramatically increases with gradual addition of ethanol. It is also worth to note that the competitive characteristic of the inhibition by edrophonium is not modified by ethanol (data not shown). Thus, our observations suggest a common effect of ethanol at the AChE active-center that brings about a decrease in the enzyme catalysis and reversible inhibition due to alteration in binding properties. 12 o
EtOH
1(]
Ki
%(v/v) m'vl
C~ O o ~
R
0.0
2.7
o
E
@
5.0
5.7
4
A
10.0
5.6
12.5
10.5
~
15.0
19.5
(3
17.5
29.3
c~
9
o ~ C 0
10
20 30 40 Edrophonium (pM)
50
Figure 3: Secondary plots of slopes of (1/V v s 1/[ATC] obtained from primary plots) v s edrophonium concentration, at various ethanol concentrations, pH 8.0, 30~ Another way to confirm the effect of the solvent on the formation and/or stability of AChE/ligand complexes is to study the reaction of the OP anticholinesterase agents that permanently block the enzyme catalysis by phosphorylating the active serine. PX well represents this family of compounds and has been chosen for our studies. AChE was incubated with various concentrations of PX in the presence of different amounts of ethanol. The residual activity of the enzyme versus the incubation time was monitored following addition of ATC and measurement of the initial hydrolytic rates. The doublereciprocal plots of the pseudo-first order inactivation rate constants (kobs) determined against the concentration of PX (figure 4) allowed us to calculate the bimolecular inhibition constant (ki) according to equation (3):
261 1
Kd
1
1
kobs- k l
k2
[PX]
k2
(3)
kl is the rate constant of solvent-induced inactivation when [PX] = 0. 100 +6 EtOH ki 10 % (v/v) M-lmin -1
80-~ 60O
4020-
0.0
4.31
@ 5.0
1.46
A
10.0
1.0
12.5
0.44
V 1 5 . 0 0.34 I I 1 2 3 4 1/[PX] (x 10-7) (M- 1) Figure 4 9Secondary plots of kobs of (residual activity vs. time) obtained from primary kinetic curves vs PX concentration at various ethanol concentrations. 0
~
v
I
0
Clearly, ethanol induces a major slope-modifying effect t h a t accounts for the d e c r e a s e in the corresponding ki values (cf. figure 4). The absence of significant variation of the intercepts at the ordinate-axis thus suggests a m a i n role of the alcohol in antagonizing the association between AChE and PX and hence, in increasing the Kd value of the complex. All our observations are consistent with a competitive modulation of the e n z y m e kinetics by ethanol c o n t r i b u t i n g to its overall effect. This solvent inhibition m a y arise from changes in the enzyme structure, thereby modifying the hydration state of the enzyme and the local dielectric constant around the active center. A decrease in the dielectric c o n s t a n t r e s u l t s in a increase of t h e electrostatic repulsion between s u b s t r a t e (or inhibitor) and its site and therefore yields lower binding. Previous studies with a-chymotrypsin also indicate a major effect of organic solvents on the dissociation constants (Km, Ki and Kd in our case) with little influence on the r a t e constants [5]. The authors proposed a t r e a t m e n t for data t h a t takes both the competitive and the dielectric component of the solvent effect into account (equation 4), which we applied to our d a t a in figure 5: K(EtOH) ~ o r K(O)
k(O) 1 1 ~ = exp(AX)withX = - ~ , k(EtOn) F.(H20- EtOn) t;(n20)
e=dielectric constant(4)
The results we obtained with three s u b s t r a t e analogues can equally be i n t e r p r e t e d as due to the cooperation of dielectric and competitive effects of ethanol t h a t alter the reaction and the inhibition of AChE at its active center.
262 := 20 # Km(EtOH)/Km(o)
~ 10
9 Ki(EtOH)/Ki(o)
0
A ki(o)/ki(EtOH)
0 0
~
1
o,5
I
I
0
5
i
I
10 15 X (x 10+2)
I
20
25
Figure 5" The linear dependence of log[Km(EtOH)/Km(0), Ki(EtOH)/Ki(0) and ki(0)/ki(EtOH)] on dielectric constant of the medium, at pH 8.0 and 30~ 3. C O N C L U S I O N Inactivation kinetics and ligand stabilization studies indicate the occurrence of conformational changes induced by ethanol at the AChE activecenter. Ligand exclusion studies further point to a main decrease of all the Km(ATC), Ki(Edro) and ki(PX) parameters induced by ethanol. This is consistent with this solvent basically affecting the substrate and ligand binding, regardless of the nature and/or charge of the molecule used. These results can quantitatively be accounted for by a combination of competitive and dielectric effects according to the following rank: PX>Edro>ATC. With regards to structural studies [6], we interpreted the effect of ethanol as due to the changes in the conformation of the amino acid residues involved in the molecular recognition of substrates and ligands.Ethanol could thus interact with aromatic amino acids required for cation-p (such as the electron-rich Y133 and W121) and/or aromatic (e.g. F330 with PX) interactions with charged or neutral compounds, respectively. The solvent effect would not be limited to the AChE active-center, since ligands selective for other sites (e.g. PAS) such as propidium or tetrabutylammonium also were shown to stabilize the enzyme against the inactivation by ethanol. Further investigations are needed to characterize the influence of the organic solvent at these sites. REFERENCES 1. Aldridge, W. N. Biochem. J., 1950, 46, 451. 2. Tanford, C. Adv. Protein Chem., 1986, 23, 122. 3. Henley, J. P. and Sadana, A. Enzyme Microb. Technol., 1985, 7, 50. 4. Ronzani, N. Analusis, 1994, 22, 249. 5. Clement G. E. and Bender, M. L. Biochemistry, 1963, vol. 2, 4, 836. 6. Ordentlich, A., Kronman C., Barak D., Stein D., Ariel N., Marcus D., Velan B., Shafferman A. J. Biol. Chem., 1993, 268,17083.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
263
Effect of a m i x e d stabilizer-denaturant system on the stability of e n z y m e activity at high temperatures : l y s o z y m e as a model
Y. Sangeeta Devi, Usha B. Nair and Rajiv Bhat* Centre for Biotechnology, Jawaharlal Nehm University, New Delhi- 110067, India Polyols viz.,sorbitol, xylitol, myo-inositol, and the disaccharide trehalose have been found to help retain the activity of lysozyme when incubated at 60eC for 24 hrs to varying extents, with myo-inositol retaining it to 72% at 0.75M concentration at pH 7.0. Guanidinium chloride, a denaturant, at 1M concentration alone retains the activity to 85% under identical conditions, while a mixture of the denaturant along with the polyols or trehalose retains the activity to ~100%. There is a good correlation of the ability of polyols to increase the transition temperature of the enzyme and the retention of biological activity. A mixture of 0.75M myo-inositol and 1M guanidinium chloride significantly prevents the activity loss even after 7 days of incubation at 60eC, suggesting that the strategy of using a cocktail mixture of moderate concentrations of a denaturant, to prevent irreversible aggregation of enzymes at high temperatures, and high concentrations of a stabilizer can be effectively used for commercial exploitation of thermolabile enzymes.
I. INTRODUCTION Enhancing the thermal stability of proteins is of prime importance for several biotechnological applications [1]. Several approaches that have been employed for this purpose include protein engineering, chemical modification of amino acids, immobilization, and solvent engineering [2-4]. Among these techniques the solvent based approach is a noninvasive one in which the covalent architecture of the protein molecule is not disturbed at all. Several solvent additives used for enhancing the thermal stability of proteins include sugars [1,5-7], polyols [8-10], amino acids [11,12], and salts [13,14]. Extensive studies on the effect of these compounds to increase the thermodynamic stability of proteins has been carried out in detail [ 15]. However, its correlation with the ability of these compounds to help retain the activity of enzymes at elevated temperatures for prolonged periods of time has not been rigorously dealt with. In addition, there are reports that although high concentrations of denaturants like guanidinium chloride (GdmCl) destabilize proteins, low concentrations can increase the thermal stability, though marginally [ 16]. * To whom correspondence should be addressed.
264 We report the results of our study on the effect of polyols viz., sorbitol, xylitol, and myo-inositol and the sugar trehalose known to increase the transition temperature (Tin) of proteins to significant extents, on the enzymatic activity of lysozyme, a model enzyme. The effect of these additives on the activity of lysozyme incubated at 60~ for 24 hrs in the presence of polyols and trehalose have been carried out. The studies have also been carried out in the presence of a mixture of 1M GdmCl and these compounds. Effect of myo-inositol, which was observed to be the best stabilizer of the activity of lysozyme, in the presence and absence of 1M GdmCl has been carried out on the long-term incubation of the enzyme up to 7 days at 60~ 2. MATERIALS AND METHODS
2.1 Materials Three times crystallized and lyophilised lysozyme (chicken egg white) from Sigma Chemical Company was used as such. The polyols sorbitol, xylitol, myo-inositol, the sugar trehalose, GdmCl, and micrococcus lysodeikticus lyophilized cells were all from Sigma and of high purity grade. The water used for making the solutions was from the Millipore Milli-Q system. Analytical grade disodium and monosodium hydrogen phosphate were procured from Merck (India). 2.2 Sample preparation and activity assay Lysozyme stock solutions at 10 Ixg/ml were prepared in 50mM phosphate buffer in the presence or absence of the cosolvent additives, using an extinction coefficient of 2.64 ml/mg.cm at 281.5 nm as reported in the literature [14]. The enzyme samples were incubated at 60~ for 24 hrs or longer periods of time after which they were incubated at 25~ for 3-4 hrs prior to the activity assay. For the activity assay micrococcus lysodeikticus cells were suspended at a concentration of 0.25 mg/ml in 50raM phosphate buffer at pH 7.0. 100~tl (11xg) of the enzyme solution was added to 2.9 ml of the cell suspension in a quartz cuvette and the decrease in absorbance was monitored at 450 nm in a Hitachi U-2000 uv-visible spectrophotometer. Change in absorbance at 450 nm per minute was calculated from which the activity of the enzyme in units/mg was determined as reported in the literature [ 17]. The activity assays were carried out at least in triplicates with the averages presented in the tables and figures. To rule out the effect of the cosolvent additives including GdmCl on the activity of lysozyme, activity assays were carried out in their presence at 25~ No change in the activity in the presence of these compounds was observed relative to the native control in the presence of buffer alone which was taken as 100%. The uncertainity in the activity measurements was within + 5%. 2.3 Thermal denaturation experiments The thermal denaturation studies of lysozyme at pH 7.0 were carried out in Shimadzu UV-160 uv-visible spectrophotometer to which a temperature programmable cuvette holder was attached which was connected to a Cecil CE-247 temperature controller module. A temperature scan of l~ was used. The wavelength selected was 301nm as reported in the literature [14]. In the presence of buffer, lysozyme at pH 7.0 got aggregated when the temperature was increased. However, in the presence of varying concentrations of GdmCl complete thermal denaturation profiles were obtained from which the Trn values could be deduced.
265 3. RESULTS Table 1 shows the data for the activity of lysozyme measured at 25~ after incubating it in the presence of various concentrations of polyols and trehalose at 60~ for 24 hrs. The data indicate that only 50% activity is retained in the absence of the cosolvent additives, while significant levels of activity are retained in their presence. Among all the cosolvents employed, it has been observed that inositol is the best stabilizer, helping retain up to 72% activity at 0.75 M concentration (which was its solubility limit under the conditions employed). It has also been observed that as the concentration of sorbitol or xylitol increases, the activity is retained to a better extent. Table 1 Activity of lysozyme at 25 ~ pH 7 after incubation at 60 ~ for 24 hours in the presence of various cosolvent additives
Table 2 Activity of lysozyme at 25 ~ pH 7 after incubation at 60~ for 24 hours in the presence of a mixture of cosolvent additives and 1M GdmCl
Cosolvent
Activity. retained
Cosolvent
Additive
(% of Nativet)
Additive
(% of Nativet)
Control
50
Control
50
Sorbitol 2M 1M 0.5M
1M GdmCl
85
63 52 45
1MSorbitol + 1MGdmCI
90
2M Sorbitol+ 1M GdmCl
98
1M Xylitol + 1M GdmCl
95
2M Xylitol + 1M GdmCl
100
0.75 Inositol+ 1M GdmCl
98
1M Trehalose+ 1M GdmCl
98
Xylitol 2M 1M 0.5M
66 55 49
Inositol 0.75M
72
Trehalose 1.5M 1M
Activity retained
72 57
t Activity of native enzyme taken as 100% Table 2 shows the effect of 1M GdmCl and a mixture of 1M GdmCl and polyols or trehalose on their ability to retain the activity of lysozyme at 60~ for 24 hrs. It is interesting to note that when 1M GdmCl is added, the activity in the presence of all the polyols and trehalose is retained to --100%, while in the presence of GdmCl alone the activity is retained to 85%. To study the effect of long-term incubation (7 days) of the enzyme at such a high
266 temperature as 60~ 0.75 M inositol was selected which was found to be the best among the series. From Fig. 1 it is evident that in the absence of the additives, no activity is retained after 7 days, while in the presence of 0.75 M inositol about 40% activity is retained which further increases to nearly 55% when 1 M GdmCl is also added to the mixture. 100. ,%. . ~ _ & ~ .
\\ ~. \~o
i - , , - ediid ........ [ -o- Q75M~ I-A- O~SM~d
~,il.
.I
+l.0MehO
~60.
~:
20"-I--____ m
o
'
~
~, Time(days)
~
8
Figure 1. Activity of lysozyme retained at 25~ pH 7.0 after incubation in the absence and presence of myo-inositol and GdmCl for several days at 60 ~
1.2-
7"5"
I
"
I
1.0 -~ 651 ~0.6
i
,
_
5s............ o.o
0.4
"
.
0.5
1.o
t
I
t
4
1.5
Odmr Cone. ( M )
2.0/
/
/___2
/---~
'
'
.11.,-3
0.2 0.0 20
"
3'0
'
4'0
'
5'0
'
6'0
Temperature (*C)
7'0
8'0
'
90
Figure 2. Thermal denaturation of lysozyme, pH 7.0 in the absence and presence of various concentrations of guanidinium chloride; 1" buffer (showing aggregation); 2: 1M GdmCl; 3: 1.5M GdmCl; 4: 2M GdmCl. Inset shows the dependence of GdmCl concentration on the transition temperature, Tm of lysozyme.
267 4. DISCUSSION
Polyhydric alcohols are known to increase the Tm of proteins by varying extents depending on the type of polyol used, e.g., sorbitol at 2 M concentration increases the Tm of lysozyme at pH 7.0 by as much as 12~ while xylitol at the same concentration increases the Tm by 8~ [18]. It has been observed that as the number of-CHOH groups present in the polyols increases, their effect on the Tm of proteins also increases. Polyols have been found to lead to the preferential hydration of proteins [ 19,20]. Recently, we have observed that the increase in the preferential hydration of proteins in the presence of polyols is due to their ability to increase the surface tension of water and that the surface tension effect is a dominant factor in modulating the solvent-mediated stability, if not the sole factor [ 18]. From the Tm studies carried out for lysozyme at different pH values earlier by us [ 18] and other coworkers [21,22], it is apparent that lysozyme denaturation is highly pH dependent. At pH 6.0 the Tm has been estimated to be nearly 74~ while the onset of transition starts at-58~ [21]. At pH 7.0, however, the denaturation is followed by sudden aggregation obscuring the calculation of Tm (Fig. 2). Since the stabilizers used push the onset of transition up by 8-12~ it is expected that lysozyme should be in the native state at 60~ in their presence. The activity assays carried out at 25~ after incubating the enzyme at 60~ for 24 hrs in the absence and presence of polyols and trehalose clearly indicate that these cosolvent additives help retain the activity of enzymes when incubated at high temperatures. It has been found out in our laboratory that inositol is a better stabilizer on a molar basis compared to sorbitol or xylitol. The activity data also indicate a similar trend. It is, therefore, obvious that there is a correlation between the ability of the cosolvent additives to increase the Tm of proteins and their effect on the retention of activity as a function of time at elevated temperatures. The increase in the retention of activity as a function of the increasing sorbitol and xylitol concentrations (Table 1) indicates that these compounds work in a nonspecific manner and that their effect is essentially water mediated. It has been observed earlier that high temperature incubation of enzymes can lead to the loss of their activity both reversibly and irreversibly depending on the temperature and time of incubation [23,24]. It is expected that at moderately high temperatures such as 60~ lysozyme may be more prone to aggregation which would lead to the loss of activity. In order to prevent the aggregation of lysozyme at this temperature, 1M GdmCl was added to the buffer solution. The Tm studies in the presence of increasing concentrations of GdmCl indicate that moderate concentrations of the denaturant, e.g., 1M do not denature the enzyme but only lower the Tm to some extent (Fig. 2). It has been reported earlier that very low concentrations of GdmCl may also increase the Tm of proteins marginally [ 16]. However, it is evident that although 1M GdmCl lowers the Tm of lysozyme by - 10~ its presence helps retain the activity to a considerable extent, indicating that 1M GdmCl is preventing aggragation of lysozyme even though it reduces its Tm and is a destabilizer. The addition of polyols or trehalose should increase the Tm and hence stabilize the enzyme in the usual way. Hence, the presence of a mixture of a mild concentration of a denaturant to prevent irreversible aggregation and high concentrations of a stabilizer work in coordination to help retain the activity of lysozyme. It is interesting to note that-100% activity of lysozyme is retained after 24 hrs of incubation at 60~ in the presence of this cocktail mixture. From Fig. 1 it is also very clear that 0.75 M inositol helps retain the activity of lysozyme to a considerable extent even after 7 days of incubation and that the addition of 1M GdmCl to the mixture increases this effect further.
268 In conclusion, the presence of a mixture of a denaturant which is at low enough a concentration not to denature the enzyme but to solubilize the aggregates or prevent aggregate formation, and stabilizers like polyols or sugars which do not bind to the protein surface but exert their influence through water mediated effects, can be used as an effective strategy to preserve the biological activity of pharmaceuticals and enzymes of commercial relevance at elevated temperatures. ACKNOWLEDGMENTS We thank Ashutosh Tiwari for assistance in the preparation of the camera ready copy of the manuscript. The financial assistance provided by Dept. of Science and Technology, Govt. of India is gratefully acknowledged. REFERENCES 1. C. Colaco, S. Sen, M. Thangavelu, S. Pinder and B. Roser, Biotechnology, 10 (1992) 1007. 2. A.R. Fersht and L. Serrano, Curt. Opin. Stmct. Biol., 3 (1993) 75. 3. E. Querol, J.A. Perez-Pons and A. Mozo-Villarias, Protein Eng., 9 (1996) 265. 4. C.O. Fagain, Biochim. Biophys. Acta., 1252 (1995) 1. 5. J.F. Back, D. Oakenfull and M.B. Smith, Biochemistry, 18 (1979) 5191. 6. J.C. Lee and S.N. Timasheff, J.Biol.Chem., 256 (1981) 7193. 7. T. Arakawa and S.N. Timasheff, Biochemistry, 21 (1982) 6536. 8. S. De Cordt, M. Hendrickx, G. Maesmans and P. Tobback, Biotechnol. Bioeng., 43 (1994) 107. 9. V. Gupta and R. Bhat, In Perspectives on Protein Engineering and Complementary Technologies, M.J. Geisow and R. Epton (eds.) Mayflower Worldwide Ltd., Birmingham, IlK p. 209 (1995). 10. C. Radha, P. Salotra, R. Bhat and R. Bhatnagar, J. Biotechnol., 50 (1996) 235. 11. M.M. Santoro, Y. Liu, S.M.A. Khan, L-X. Hou and D.W. Bolen, Biochemistry, 3 (1992) 5278. 12. S. Gopal and J.C. Ahluwalia, J. Chem. Soc. Faraday Trans., 89 (1993) 2769. 13. T. Arakawa and S.N. Timasheff, Biochemistry, 21 (1982) 6545. 14. T. Arakawa, R. Bhat and S.N. Timasheff, Biochemistry, 29 (1990) 1914. 15. S.N. Timasheff, In Protein-Solvent Interactions, R. B. Gregory (ed.), Marcel Dekker, Inc., New York, p. 445 (1995). 16. L.M. Mayr and F. X. Schmid, Biochemistry, 32 (1993) 7994. 17. D. Shugar, Biochim. Biophys. Acta., 8 (1952) 302. 18. J.K. Kaushik, Ph.D. thesis, Jawaharlal Nehru University, New Delhi (1996). 19. K. Gekko and T. Morikawa, J. B iochem. (Tokyo), 90 (1981) 39. 20. G. Xie and S. N. Timasheff, Protein Sci., 6 (1997) 211. 21. T. Arakawa and S. N. Timasheff, Biophys. J., 47 (1995) 411. 22. P.L. Privalov and S. A. Potekhin, Methods Enzymol., 131 (1986) 4. 23. T.J. Ahem and A. M. Klibanov, Science, 228 (1985) 1280. 24. D.B. Volkin and R. Middaugh, In Stability of Protein Pharmaceuticals, Part A: Chemical and physical pathways of protein degradation, T. J. Ahem and M. C. Manning (eds.), Plenum Press, New York, p. 215 (1992).
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. HaUing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
269
A simple folding method for high level production of the hydrophobic disulfide bonded hepatitis B X protein by inclusion body route and its structural analysis I. Marczinovits a, J. Molnhra, M. Z. Keleb, P. T. Szab6b and T. Janhkyb Departments of aMicrobiology and bMedieal Chemistry of Albert Szent-Gyorgyi Medical University, P.O. Box 8, 6720 Szeged, Hungary*
ABSTRACT Insoluble inclusion bodies are frequently formed upon recombinant protein expression in transformed micro-organisms. As these inclusion bodies contain the recombinant protein in a highly enriched form, it is likely that the inclusion body route for production of recombinant proteins will remain commercially important. However, following inclusion body strategy, the main obstacle is to regain the native, active protein structure in considerable quantity. New folding procedures have been developed for efficient in vitro reconstitution of complex hydrophobic, multidomain, highly disulfide-bonded proteins. Nevertheless, despite numerous experimental data in this field, protein folding is an empirical science and optimal procedure has to be determined on a case-by-case base. Here we demonstrate a solubilization and refolding protocol for the GST-X (glutathion Stransferase-I-IBVx) fusion recombinant protein. With rapid dilution of the protein solution into simple folding conditions (2.66 M urea, pH: 6.5, 4~ air oxidation) renaturation process was carried out at 2-3 mg/ml protein concentration. The proper conformation of the recombinant protein was supported by complete separation of the fusion partners by X a protease. Localization of the disulfide bonds between cysteine residues in the refolded HBx protein was established by electrospray mass spectrometry (ESI-MS) combined with high performance liquid chromatography (HPLC). The disulfide linkages were between Cys3 and Cys 137 , Cys61 and Cys69, Cys 17 and Cys 143 while Cys 115 was flee.
INTRODUCTION Hepatitis B virus (HBV) is an important virus because it can cause both acute and chronic hepatitis and has been linked to the formation of primary hepatoceUular carcinoma [1]. The X gene is the smallest of the four deduced open reading flames (ORFs) of HBV DNA and has the potencial to encode a 154 amino acid polypeptide with hydrophobic and disulfide-bonded characters. This protein may reveal diagnostic significance as there is connection between the This work was supportedby the Hungarian National Committeefor Technical Development(OMFB) No. 939748-0543 and by the National ScientificResearch Foundation(OTKA) T 026624.
270 progression from the chronic to malignant state of the disease and the expression of the X protein in infected hepatocytes [2]. To produce the X protein in reasonable quantity we have used the high expression vector system pGEX (Pharmaeia) and large quantity of hepatitis B antigen (HBxAg) was prepared from inclusion body fraction of the E. coli cell lysine. In the present paper an optimal in vitro refolding protocol for production of HBx protein is presented. As the X protein is a complex protein, to gain insight into the structural properties of the protein: how disulphide bonds form upon recreation of native conformation, positions of disulfide bonded cisterns were examined by electrospray mass spectrometric measurements that resulted in linkages between Cys3 and Cys 137 , Cys61 and Cys69, Cys 17 and Cys 143. MATERIALS AND METHODS
Plasmodia construction, expression and purification of GST-X fusion protein, cleavage and separation of the fusion partners were as described earlier [3] and described here briefly. The truncated HBx gene was cloned through transient expression vectors into the fusionexpression vector pGEX-3X (Pharmacia), producing the pGEX-3XXBF recombinant plasmid. According to the cloning strategy, the X protein sequence was truncated by 9 and 11 amino acids and flanked by 6 and 4 amino acids at the N- and C-terminals, respectively. E. coli DH5a cells harbouring the fusion-expression plasmodia construction was grown in LB medium and induced with 0.5 mm isopropyl b-D-thiogalactopyranoside (IPTG) for 3 hours. Bacteria were harvested by eentrifugation then treated with lysozyme and disrupted by sonication. The soluble and insoluble fractions of the E. coli lysate were separated by eentrifugation. The bulk of the GST-HBxAg fusion protein was selectively solubilized from the inclusion bodies by vortexing for 3-4 min in 8 M urea prepared with deionized distilled water at pH 6.5. The extracted GST-X protein was renatured by diluting the protein solution at 6-9 mg/ml in 8 M urea with two volumes of deionized distilled water. In the refolding step, disulfide bonds were formed by air oxidation during the storage of the protein solution in 2.66 M urea for one day at 4~ The renatured fusion protein was digested with with Xa protease in vitro at 2-3 mg/ml at a substrate to enzyme ratio of 50:2 (w/w) for 3 hours. The cleavage reaction was carried out in deionized distilled water containing 2.66 M urea at pH 6.5 and 25~ The cleaved fusion partners were further separated on FPLC Superdex 75 column (Pharmacia). Purification and separation procedures were analysed by electrophoresis in 15 % SDS-polyacrylamide gel (SDSPAG).
Identification of disulfide bonds: Following protease cleavage, the GST-HBx fusion protein was reacted with vinyl-pyridine to alkylate free cysteine(s) and separated in native SDSTricine-PAG. Band with 16 kDa molecular mass was cut from and digested in the gel by trypsin [4]. The digested fragments were eluted from the gel, separated and analysed by capillary reversed phase HPLC coupled to the mass spectrometer (Finnigan TSQ-7000, Finnigan MAT, US). The I-IPLC system contained ABI 140C syringe pump and 785A UV detector (ABI, US). Elution was performed with a 70-min linear gradient from 5-90% solvent B (0.04 % TFA in 80 % aqueous acetonitrile) in solvent A (0.05% aqueous TFA) on reversed phase column [32x250 mm, C-18 (300 A, 5 l.tm), LC-Packings, Netherlands]. A theoretical tryptic digestion
271 was made by a computer program where all the possible alkylated and disulfide bonded cysteines were generated. These date were compared with the molecular weights of peptide fragments found by the HPLC-ESI-MS analysis [5]. RESULTS AND DISCUSSION Purification and renaturation of GST-X fusion protein More than 80% of the GST-X fusion protein was accumulated in inclusion bodies. Taking into account that protein folding processes are not strongly affected by other proteins in the renaturation environment, rigorous purification of the inclusion body protein may not be a prerequisite for efficient refolding. Therefore, we followed a quick selective solubilization procedure using deionized distilled water-at pH 6.5 in the presence of 8 M urea. Upon a short time of extraction by vortexing high purity and large quantity of fusion protein were obtained (Fig. 1A, lane 2). Selectivity of the extraction and solubility of the dissolved fusion protein could only be obtained at pH 6.5. Increasing the pH of the extraction buffer to pH 8.0 resulted in not only considerable impurities derived from the E. coli (not shown) but greatly enhanced tendency to precipitate of the extracted protein.
Fig. 1. Digestion of the GST-X protein with factor Xa after refolding (A) and analysis of the gel filtration process by electrophoresis in SDS-PAG (B). (A) Lane 2: selectively solubilized and refolded fusion protein. Lane 1: proteins of enzymic reaction after 3 h digestion. Reaction conditions were as described in materials and methods. Lane 3: molecular mass standards in kDa. The position of the fusion protein (fp), glutathione S-transferase (GST), and the pX equivalent are indicated. (B) Lane 1: protein sample before loading onto the Superdex TM 75 HR FPLC column; lanes 2-8 represent proteins from the peak fractions. Lane 9: marker proteins in kDa. Electrophoresis was performed on 15% SDS-PAG, and stained with Coomassie Brillant blue.
272 The latter may be due to the high isoelectric point (I=8.5) of the X protein. Although the structural characteristics of the X protein (52% of the amino acids in the protein is hydrophobic and the protein contains disulfide bonds) may also be a potential reason. As a result of the selective solubilization, 10-15 mg of fusion protein can be obtained from 100 ml of culture. Refolding of solubilized inclusion body proteins is usually performed by dilution or dialysis. Prediction for the preference of refolding of a particular protein by dilution or by dialysis can not be given. Both approaches can be beneficial as well as detrimental for correct folding. Therefore, the optimum procedure has to be found and is protein-specific. In our case, rapid dilution of the denaturant proved to be beneficial for renaturation of the inclusion body protein without any precipitation at relatively high concentration (2-3 mg/ml). In contrast, gradually removal of the denaturant by dialysis induced quantitative precipitation of the protein and resuited in a soluble form of that up to 200 ~tg/ml only. The above phenomenon may be the consequence of the population of folding intermediates at an intermediate denaturant concenration which induces quantitative precipitation. Following cleavage of the fusion partners the X protein contains seven cysteine residues. Therefore, having formed a compact conformation under the particular folding technique we addressed the question at what variation of covalent disulfide bonds were generated. The exact positions of disulfide bonds could be given by mass spectrometry measurement.
Cleavage and separation of the refolded GST-X fusion protein, respectively The advantageous conformation of the GST-X fusion protein was reflected by the fact that the enzymic reaction by Xa protease was akeady complete after a 3-h incubation period at 25 ~ (Fig. 1A, lane 1). After proteolytic cleavage the GST and the viral partner were separated on a Superdex 75 FPLC column. The peak fractions were analysed for their purity by means of SDS-PAG electrophoresis (Fig. 1B). Pure fractions of the X protein were collected and stored at -20~ Identification of disulfide bonds Two alkylated fragments with molecular weights of 792.2 and 1985.8 were found (Fig. 2, a and b; Fig3, A and B ). These fragments were missing from the LC-MS chromatograms of
Fig. 2. Mass chromatograms of disulfide bonded peptides. The figure contains LC-MS data. The a-f traces are selected ion chromatograms of specific ions where peaks are related to sample fractions.
273 the tryptie digestion of the nonalkylated protein (not shown) indicating that only the cysteine at position 115 is free and the other 6 ones are disulfide bonded. Two fragments with molecular weights of 1535.9 and 2164.9 indicate that the eysteines at position 3 and 137 are connected (Fig 2, c and d; Fig. 3, C and D). The second fragment originated from incomplete digestion of X protein by trypsin as identified by multiple charged ions. Two peaks have been found in the mass chromatogram of the ion of 1082.3 (Fig. 2, d). The first one is the doubly charged molecular ion of the disulfide bonded peptides, the second
Fig. 3. ESI mass spectra of identified cysteine-containing peptide fragments
274 one is related to the auto digestion of trypsin (Fig. 2, d). Two peaks were found in the mass chromatogram of ion m/z=774.1 (Fig. 2, e). This species might be the doubly charged ion of the disulfide bonded peptide fragment 61-69 having the molecular weight of 1546.9 (Fig. 3, E). In order to determine which peptide fragment contains the disulfide bond, both peptides were reduced and pyridylethylated. The mass spectra of non-alkylated and alkylated peptides indicate that the first peak contains the peptide with disulfide bridge between cysteines at positions 61 and 69 (Fig. 3, F). The other peptide fragment originates from the autodigestion of trypsin. The only possibility for the third disulfide bridge is the disulfide bond between the cysteines of 17 and 143. The molecular weight of a tryptic fragment with this connection is 2090.3 (Fig, 3, (3). This molecule has been found and can be seen in the mass chromatoram of ion 1046.6 (the doubly charged molecular ion) (Fig. 2, f). Not only the doubly but the triply charged ion of this fragment can also be detected in the mass spectrum (Fig. 3, G). On the bases of these data it can be concluded that the Cys 115 is free, and three disulfide bonds can be found between Cys3*-Cys137; Cys61-Cys69 and Cysl7-Cys ]43 (Fig. 4). .A unique arrangement of disulfide bonds of HBx expressed in E. coli as inclusion body protein was determined in [6] which differ from our findings. That may be due to the differences of the two viral DNA and protein sequences. 3* 17 G~STDPARDVL~:,RPVGAESRGRPFSGSLGTLSSPSPSAVSTDHGAHL 61
69
SLRGLPV~FSSAGP~LRFTSARRMETTVKAQPFLPKVLHKRTLGLSV 115
137
MSTTDLEAYF~FKDWEELGEEI~KVFVLG~KL~NSS ,
,
,
~
143 I
Fig. 4. The amino acid sequence of the cloned HBx protein indicating the positions of the disulfide bonds. Bold letters represent the amino acid residues of the X protein flanking with sequences derived from the cloning vectors. 3* indicates cysteine residue has been originated from the transient cloning vector. Here we report an optimal and convenient refolding method for a rather hydrophobic, and disulfide bonded viral protein, HBx. To characterize the outcome of the oxidative refolding process we have applied mass spectrometry and determined positions of disulfide bonds within the HBx protein. REFERENCES 1. W.S. Robinson, Ann. Rev. Med., 45 (1994)297. 2. Q. Su, Y.F. Liu, J.F. Zhang, S.X. Zhang, D.F. Li, J.J. Yang, Hepatolo~, 20 (1994) 788. 3. I. Marczinovits, Cs. Somogyi, A. Patthy, P. N~meth and J. Moln/~r, J. Biotechnol., 56 (1997) 81. 4. T.D. Lee and J.E. Shively, Methods in Enzimology, 193 (1990) 361. 5. S.D. Patterson and R. Aebersold, Electrophoresis, 16 (1995) 1791. 6. A. Gupta, T.K. Mal, N. Jayasuryan an V.S. Chauhan, Biochem. Biophys. Res. Commun., 212 (1995) 919.
Protein engineering and Thermophile enzymes
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
RIGIDITY
OF THERMOPHILIC
277
ENZYMES
Angelo Fontana, Vincenzo De Filippis, Patrizia Polverino de Laureto, Elena Scaramella and Marcello Zambonin CRIBI Biotechnology Centre, University of Padua, via Trieste 75, 35121 Padua, Italy
Enzymes and proteins isolated from thermophilic microorganisms are not only unusually stable to heat and protein denaturants, but also display enhanced protein rigidity in respect to that of their mesophilic counterparts. The molecular rigidity of thermophilic enzymes appears to explain why their specific activity at room temperature often is less than that of the corresponding mesophilic enzymes, considering that an appropriate level of protein mobility is required for catalysis. Evidence of protein rigidity can be obtained from hydrogen exchange measurements, molecular dynamics simulations, by computing flexibility indices based on crystallographic data, as well as by proteolysis experiments. Although the structural and functional complexity of proteins likely does not allow firm generalizations, it can be proposed that thermophilic enzymes are rigid molecules, but not optimally active at ambient temperature. Considering that extremophiles appeared earlier on hearth in a hotter environment, it can be suggested that present-day mesophilic enzymes evolved to be more flexible, and thus more labile, in order to optimize their catalytic function.
1. INTRODUCTION The structural, functional and stability properties of enzymes isolated from thermophilic bacteria have attracted the interest of many investigators in the last two decades (Perutz & Raidt, 1975; Jaenicke, 1981, 1991, 1996; Fontana, 1990; Adams, 1993; Gupta, 1993; Vieille & Zeikus, 1996; Vieille et al., 1996; Arnold, 1998). Thermophilic enzymes possess unusual stability towards the denaturing action of heat and protein denaturants and therefore can be used as biocatalysts under rather harsh environmental conditions (Sonnleitner & Fiechter, 1983; Zamost et al., 1991; Adams et al., 1995; Ffig~in, 1995). Moreover, the unusual properties of thermophilic enzymes and proteins prompted their use as suitable protein models for addressing a number of fundamental problems in current protein research (Jaenicke, 1991, 1996). The aims of studies on thermophilic enzymes complement those of modem protein engineering studies by genetic methods, since both studies can ultimately lead to a better and, hopefully, quantitative understanding of the structure-stability-function correlations in
278 proteins (Mattlaews, 1991, 1993; Fontana, 1991). Indeed, some useful guidelines for enhancing protein thermostability have been deduced from studies on thermophilic enzymes and successfully applied in protein research (Fersht & Serrano, 1993; Men6ndez-Arias & Argos, 1989; Russell & Taylor, 1995; Fhghin, 1995). The numerous studies carried out in the past on the molecular properties of thermophilic proteins revealed that quite subtle structural differences between a thermophilic and a mesophilic protein are sufficient to cause the observed unusual stability (Jaenicke, 1981, 1991, 1996; Fontana, 1990). In particular, the analysis of homologous proteins from thermophilic and mesophilic sources in terms of amino acid sequences and three-dimensional structures (Argos et al., 1979; Men6ndez-Arias & Argos, 1989; Vogt et al., 1997; Vogt & Argos, 1997) revealed that the enhanced thermostability cannot be attributed to a common determinant, but is the result of a variety of stabilizing effects brought about by hydrophobic interactions, ionic and hydrogen bonding, disulfide bonds, metal binding, and so on. Moreover, the effects of amino acid exchanges in a protein can be cumulative (Wells, 1990), so that few amino acid substitutions, giving each an extra free energy of stabilization of few kcal/mole, can lead to a cumulative effect of significant enhancement of protein stability (Matthews, 1991, 1993; Zhang et al., 1995). Nowadays there is a general consensus that a universal molecular mechanism of thermostability cannot be proposed, since different proteins may be stabilized in different ways. Nevertheless, in a recent study (Vogt et al., 1997), the stabilizing interactions and forces occurring in 16 families of proteins of known three-dimensional structure and different thermal stability have been carefully analyzed. It was concluded that an increased number of hydrogen bonds and salt bridges appears to play a major role in enhancing protein thermostability. The variety of physical and chemical reasons that have been advanced in order to explain the enhanced thermostability of thermophilic enzymes have been recently reviewed (Querol et al., 1996; Vogt et al., 1997). Why all enzymes in nature have not yet evolved to be more stable, despite the three billion years of evolution? Since this has not occurred, there should be good reasons for it. The hypothesis can be advanced that an enzymatic apparatus constituted by extremely stable and thus rigid enzymes is not required for an organism living in a normal habitat, while thermolabile enzymes are more suited for life under common physiological conditions. Indeed, protein molecules and enzymes exploit their functions via their dynamic nature characterized by backbone and side-chain mobility (Gurd & Rothgeb, 1979; Alber et al., 1983; Fersht, 1985; Kraut, 1988), as well as domain motion (Huber, 1979), and exceedingly stable proteins are therefore too rigid molecules to be optimally functioning (Zuber, 1981; Hochachka & Somero, 1984; Jaenicke et al., 1996; Fontana, 1990, 1997). In this article we will summarize experimental findings that indicate that the stable thermophilic enzymes are quite rigid molecules at room temperature with respect to their mesophilic counterparts, as a direct consequence of the "clamping" effect of protein structure brought about by the interactions and forces which stabilize thermophilic proteins. No attempt is made here to provide a complete coverage of the
279
. Figure 1 9 Effect of temperature on the catalytic activity of enolase from rabbit, yeast, Thermus aquaticus and Thermus X1. All assay solutions contained 2 mM 2phosphoglycerate and 1 mM MgSO4. The assay solution was placed in a cuvette and brought to the appropriate temperature in a thermostatted cell holder of the spectrophotometer. The enzymatic reaction was initiated by adding an aliquot of the enzyme solution. Data taken from Stellwagen et al. (1973), Barnes & Stellwagen (1973) and Stellwagen & Barnes (1976).
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Temperature (~ vast literature dealing with structure, stability, rigidity and functional properties of thermophilic enzymes, so that the reader may fmd herewith some personal selections and omissions of issues as well. In particular, data will be presented which are in line with the proposed stability-rigidity correlation in thermophilic enzymes, even if we are aware that the complexity of protein molecules does not permit to affirm the strict generality of this correlation. Despite the existence of numerous reviews, monographs and books dealing with the molecular and stability properties of the enzymes isolated from extremophiles, an account which specifically addresses and discusses the rigidity of thermophilic enzymes is missing. The authors therefore hope that their effort will be useful and will prompt new interpretations and ideas, as well as fiu~er experimentation. 2. INVERSE CORRELATION BETWEEN PROTEIN STABILITY AND CATALYSIS Thermostable enzymes isolated from thermophiles are expected to be rigid molecules at room temperature and consequently this rigidity should have an adverse effect on their catalytic efficiency (Zuber, 1981), since it is known that an appropriate degree of flexibility is required for enzyme catalysis (Alber et al., 1983; Fersht, 1985; Kraut, 1988). In fact, thermophilic enzymes are usually poor catalysts at room temperature and assays of their enzymatic activity should be conducted at a temperature close to that of optimum temperature growth of the organism from which the etm3nne has been isolated. Often it has been observed that at moderate temperature the specific
280 activity of a thermophilic enzyme is less than that of its mesophilic counterpart and only at high temperature is sufficiently flexible to be fully active and yet rigid enough not to be denatured (Jaenicke, 1981; Fontana, 1990). The inverse correlation between enzyme activity and thermostability has been demonstrated in several cases. As shown in Fig. 1, the specific activity (i.e., catalytic efficiency) at 37~ of enolase from rabbit, yeast, Thermus X-1 and Thermus aquaticus YT-1 at room temperature is of the inverse order of their thermostability (Stellwagen et al., 1973; Barnes & Stellwagen, 1973; Stellwagen & Barnes, 1976). At high temperature, thermophilic enolases acquire enhanced mobility and catalytic potency, while the mesophilic ones do not resist to the denaturing action of heat. The relationships between stability, dynamics and activity in 3-phosphoglycerate kinase (PGK) from yeast, the moderate thermophile B. stearothermophilus and the extreme thermophile Thermus thermophilus HB8 have been carefully analyzed by Varley & Pain (1991). This enzyme is a two domain protein operating through a hinge bending mechanism and consequently its activity would be expected to be dependent upon large-scale dynamics due to domain movements (Banks et al., 1979). The thermophilic PGK is more stable than the yeast enzyme, but less active at 25~ The maximum specific activity of PGK from the three sources reached similar values at the temperature of optimal growth of the organism from which the enzymes are derived. The conclusion of this study was that a proper balance and correlation exist between thermodynamic stability, dynamics and specific activity and, in particular, that increased stability constrains conformational dynamics and enzyme activity (Varley & Pain, 1991). Several other proteins of the same function from mesophilic and thermophilic bacteria exhibit similar levels of activity at the temperature of optimal growth of the organism from which they are derived (Zuber, 1981; Hochachka & Somero, 1984; Somero, 1995; Vihinen, 1987; Rees & Adams, 1995; Jaenicke et al., 1996). Enhancement of protein stability by site-directed mutagenesis is a major aim of modem protein engineering studies (Alber, 1989; Matthews, 1991, 1993). The proposal that stability/rigidity hampers enzyme catalysis, and protein function in general, predicts that enhanced protein stability would be achieved at the expenses of catalytic potency. Indeed, the inverse correlation between stability and catalysis was documented by measuring stability-activity relationships in four mutants of kanamycin nucleotidyltransferase with single or double amino acid replacement(s) (Matsumura et al., 1986). Matthews and coworkers (Zhang et al., 1995) prepared a series of mutants of T4 lysozyme and demonstrated that the combination of several point mutations in the protein leads to mutant species with an additive increase in thermal stability, but also to a gradual decrease in activity. Moreover, the hypothesis that amino acid residues at the active site of an enzyme are mobile and thus not optimized for protein stability has been tested by analyzing the effects of amino acid replacements at the active site of T4 lysozyme (Shoichet et al., 1995). It has been demonstrated that it is possible to produce active site mutants with improved stability but reduced activity. A variety of protein engineering experiments designed to increase protein stability also resulted the production of more stable mutants at the expenses of catalysis or of other functional
281
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~ . Salt-mediated activation of thermolysin. The enzymatic assays were conducted at 25~ in Tris buffer, pH 7.5, in the presence of the indicated concentrations of NaBr with furylacryloyl-glycyl-L-leucine amide (FAGLA) ( 9 or benzoyl-glycylphenylalanyl-alanine (O). Data taken from Holmquist & Vallee (1976).
Figure 3. Activation by KCI of isocitrate lyase from Bacillus stearothermophilus BS1. The enzymatic assays were conducted under standard conditions in the presence of various
concentrations (0-0.4 M) of KC1 at 22~ (0), 30~ (O), 40~ (!1), 45~ (El), 50~ (A) and 55~ (zx). Data taken from Griffiths & Sundaram (1973).
properties of the proteins (see Shoichet et al., 1995, for a list of these experiments). Of interest, Kidokoro et al. (1995) prepared a series of mutants of thermolysin by replacing the Gin residue in position 119 of the polypeptide chain. All mutants were less thermostable than the wild-type species, but displayed a much improved enzyme activity, in line with the proposed correlation between enzyme lability/flexibility and catalysis. 3. ENZYME ACTIVATION AT LOW DENATURANT CONCENTRATION Thermophilic enzymes can be activated in the presence of protein denaturants (salts, organic solvents, urea, guanidine hydrochloride) at moderate concentrations. For example, glyceraldehyde-3-phosphate dehydrogenase from Thermus thermophilus is activated when salts and ethanol are added to the enzymatic assay solution (Fujita et al.,
282 1976). 6-Phosphogluconate dehydrogenase from B. stearothermophilus is more active in the presence of organic solvents (dioxane, dimethylformamide, acetone) (Veronese et al., 1984). Similarly, a number of organic solvents have been shown to substantially activate the malic enzyme from the extreme thermoacidophilic archaebacterium Sulfolobus solfataricus (Guagliardi et al., 1989). The catalytic activity of malate dehydrogenase isolated from several thermophilic bacteria is strongly activated, if the assays are performed in the presence of 3-20% acetone, 4-8 M urea or 0.5-2 M guanidine hydrochloride (Sundaram et al., 1980). Triosephosphate isomerase from Thermotoga maritima is activated (up to 180%) when assayed in the presence of 2 M guanidine hydrochloride (Beaucamp et al., 1997). Fig. 2 and 3 illustrate the salt-mediated activation effects for thermolysin (Holmquist & Vallee, 1976) and isocitrate lyase from B. stearothermophilus BS1 (Griffiths & Sundaram, 1973), respectively. It is seen that thermolysin can be activated up to about 25-fold and 5-fold if assayed in the presence of 5 M NaBr with furylacryloyl-Gly-Leu-NH2 or benzoyl-Gly-Phe-Ala as substrate, respectively (see Fig. 2). The data of Fig. 3 indicate that the KCl-mediated activation of thermophilic isocitrate lyase occurs at moderate temperature only, while at 55~ the salt activation is marginal. These data can be interpreted as indicating that at relatively high temperature the thermophilic enzyme becomes flexible enough to be optimally active, while at 2230~ some salt-mediated weakening of the interaction and forces leading to its rigid structure is required for improving its catalytic potency. These various observations can be taken as an indication that thermophilic enzymes are activated by some loosening of their structure in the presence of protein perturbants. The denaturant-mediated enhancement of catalysis can result from a very limited change of structure/dynamics of the enzyme. Far-UV circular dichroism or fluorescence emission measurements do not detect changes in secondary and tertiary structure, respectively, with activated Thermotoga triosephosphate isomerase dissolved in the presence of 2 M guanidine hydrochloride (Beaucamp et al., 1997). 4. ANALYSIS OF PROTEIN RIGIDITY/FLEXIBILITY The rigidity of thermophilic proteins has been verified utilizing hydrogen exchange measurements (Wraba et al., 1990; Rehaber & Jaenicke, 1992), fluorescence quenching (Varley & Pain, 1991) and theoretical calculations (Vihinen, 1987; Vihinen et al., 1994; Lazaridis et al., 1997). The H-D exchange rates in globular proteins are a reflection of protein mobility/flexibility (Delepierre et al., 1983), as documented for example by the inverse correlation between the melting temperature (Tin) and H-D exchange rates in a series of derivatives of basic pancreatic trypsin inhibitor (Wiithrich & Wagner, 1979; Wiithrich et al., 1980). The higher the exchange rate, and thus the higher the flexibility, the lower is the denaturation temperature. Elongation factor Tu of Thermus thermophilus shows a reduced rate of exchange in respect to the more thermolabile E. coli factor (Tsuboi et al., 1978).
283 2.0
Figure 4. Fluctuations along the polypeptide chain of rubredoxin from Pyrococcus furiosus (RdPf) (solid line) and from Desulfovibrio vulgaris (RdDv) (dashed line). Molecular dynamics simulations were performed at 300 K and the simulation extended to 400 ps. Data taken from Lazaridis et al. (1997).
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Since the rate of quenching of tryptophan fluorescence in a globular protein is dependent upon the fluctuation of protein structure (Eftink & Ghiron, 1976, 1981), Varley & Pain (1991) measured the ability of acrylamide to quench the fluorescence of a buffed tryptophan in thermophilic and mesophilic 3-phosphoglycerate kinase. At 25~ the rate of quenching was more than one order of magnitude less in the Thermus than in the yeast enzyme. The increased rigidity of thermophilic proteins in respect to their mesophilic counterparts was also documented by calculating protein flexibility indices (F) derived from normalized B-values (temperature factors determined crystallographically) of individual amino acids in several refined three-dimensional structures of globular proteins. The results of these calculations showed that F-values correlate with protein stability, i.e., that rigidity correlates with thermostability of proteins (Vihinen, 1987; Vihinen et al., 1994). More recently, a comparative analysis of the molecular rigidity of a thermophilic and a mesophilic protein has been conducted utilizing molecular dynamics simulations (Lazaridis et al., 1997). At room temperature, the fluctuations along the polypeptide chain of rubredoxin from the hyperthermophilic archaeon Pyrococcus furiosus (RdPf) are somewhat reduced in respect to those of the homologous rubredoxin from the mesophile Desulfovibno vulgaris (RdDv) (see Fig. 4). The average value of fluctuation for all residues is 0.8 A for RdPf and 0.9 A for RdDv. Residues 2-25 show similar fluctuations in the two proteins, while residues 25-50 exhibit higher fluctuations in the mesophilic protein. 5. PROBING OVERALL AND LOCAL PROTEIN FLEXIBILITY BY PROTEOLYSIS The overall rigidity of the stable proteins from thermophilic microorganisms can be evidenced by the use of proteolytic enzymes as probes of protein structure and
284 dynamics (Fontana et al., 1993). In analogy to all enzymatic reactions, the proteolytic cleavage of a polypeptide chain occurs only if the site of cleavage can bind and adapt itself in a specific way to the stereochemistry of the active site of the protease. This is difficult to achieve with native globular proteins, whereas denatured/unfolded proteins are much more susceptible to proteolysis. On this basis, it is expected that only the unfolded species of a globular protein is attacked by a protease and that the actual equilibrium between the native (N) and denatured (D) form of protein is controlling the rate of protein degradation. The N<---~Dequilibrium is expected to be shifted towards the N form under physiological conditions with the stable thermophilic proteins, so that proteolysis would be hampered. The correlation between protein stability/rigidity and resistance to proteolysis is also substantiated by the fact that proteins with relatively short half-lives in vivo have high protease susceptibility and are generally unstable in vitro (Goldberg & Dice, 1974; McLendon & Radany, 1978). Unusual resistance to proteolysis of thermophilic enzymes has been documented in a number of cases, thus verifying their overall rigidity. 6-Phosphogluconate dehydrogenase from B. stearothermophilus is much more resistant at room temperature to proteolysis than the same enzyme from yeast (Veronese et al., 1984). Similarly, the activity loss due to proteolysis of asparaginase and 13-galactosidase from thermophilic and mesophilic sources correlates with the thermal instability of the enzymes (Daniel et al., 1982). Oligo-l,6-glucosidase from B. thermoglucosidwus was more resistant against proteolysis than its homologous counterpart from B. cereus (Suzuki & Imai, 1982). An inverse correlation between protein stability (and thus rigidity) and proteolytic degradation was reported for mutants of the ct-subunit of tryptophan synthetase from E. coli (Ogasahara et al., 1985). Similar observations were reported for mutants of kanamycin nucleotidyl transferase (Matsumura et al., 1986) and T4 lysozyme (Schellman, 1986). In a recent study, proteolysis experiments were employed to monitor the stability/rigidity of mutants of neutral protease from B. stearothermophilus prepared by introducing Pro residues at the level of the active site helix of the enzyme (Nakamura et al., 1997). As shown in Fig. 5, there is an inverse correlation between protein thermal stability, and thus rigidity, and rate of chymotryptic hydrolysis of the Xaa~Pro mutants. A systematic study of the behaviour of thermophilic enzymes towards proteolysis was conducted using the metaUoendopeptidase thermolysin as a model protein (Fontana et al., 1986). In this case, specific experimental conditions of limited proteolysis by added protease or autolysis of thermolysin permitted the isolation and characterization of several "nicked" thermolysin species constituted by two as well as three fragments associated in stable complexes. The pattern of limited protein fragmentation of thermolysin allowed to infer molecular aspects of the protein-protein recognition process underlying the proteolytic event and, in particular, to establish that flexible loops of the thermolysin molecule are the most susceptible sites of proteolysis. Of note, a striking correlation was shown to exist between sites of proteolysis and sites of high segmental mobility determined crystallographically (B-values). The conclusion was
285
Figure 5. Inverse correlation between 76 I I I I i thermostability and susceptibility to proteolysis of wild-type ( 9 mutant (o) 74 ~ ' ~ O I1 B. stearothermophilus neutral protease. 40P I \ The protease was dissolved at 37~ in 50 721-mM Tris-HC1 buffer, pH 7.5, containing 5 ~ [ ON~141P mM CaCI2 and 3 mM phosphoramidon and v then heated at 70~ for 10 min. Under ~ 70 ~ ~_ these solvent conditions the B. ~ D153P(~Wild-type stearothermophilus protease is inactivated ~ 68 by removal of its functional zinc ion, while E the stabilizing calcium ions remain bound to the protein. Proteolytic digestion was ~ 6 6 conducted at 37~ by adding ~chymotrypsin at an enzyme to substrate 64ratio of 1:10 (by weight). The relative rate of initial hydrolysis was determined by 62 quantitation of the free amino groups formed during digestion. The thermostability of wild-type and mutant 60 I I I ! I neutral protease was evaluated from the 0 1 2 3 4 5 6 irreversible thermal denaturation curves Relative Rate of Hydrolysis from 20 to 90~ of the circular dichroism signal at 222 nm versus temperature. From these curves the melting temperature (Tin) was calculated. The single site mutants of neutral protease (I140P, D141P, L147P and D153P) are indicated in the figure, while the number refers to the site of the polypeptide chain where the Xaa~Pro exchange has been introduced. Data taken from Nakamura et al. (1997). reached that a mechanism of local unfolding dictates the phenomenon of limited proteolysis of globular proteins (Fontana et al., 1986). On this basis, a procedure was proposed for probing the sites of flexibility of a protein by proteolysis experiments (Signor et al., 1990), as well as for stabilizing proteins against proteolytic degradation by site-directed mutagenesis of the protein sites most prone to proteolysis (Braxton & Wells, 1992; Fontana et al., 1993; Ros6 et al., 1993; Van den Burg et al., 1998a). 6. ENGINEERING PROTEIN STABILITY/RIGIDITY The advent of site-directed mutagenesis techniques made possible in recent years to produce a large variety of protein mutants and, in particular, to analyze the forces and interactions thought to be important in determining protein stability (Alber, 1989; Matthews, 1991, 1993). These numerous studies allowed a detailed, and often quantitative, analysis of the contributions to protein stability of hydrogen bonds, electrostatic interactions, hydrophobic effects, disulfide bridges, ion binding and so on (Dill, 1990). As a result of these intensive and systematic investigations, some strategies
286 for improving protein stability by the use of genetic methods have been developed. As a matter of fact, the successful production of engineered variants of proteins that are more stable than the wild-type protein has been often described (Matthews et al., 1987; Matthews, 1991, 1993; Matsumura et al., 1986, 1989; Fontana, 1991; Zhang et al., 1995; Van den Burg et al., 1998b). The results of protein engineering experiments complement those derived from studies on thermophilic enzymes, since both approaches aim to unravel structure-stability relationships in proteins (Fontana, 1991; Fersht & Serrano, 1993; Russell & Taylor, 1995). Indeed, some strategies of protein stabilization developed by protein engineers parallel those followed by nature in engineering by evolution and natural selection the stable enzymes from thermophilic microorganisms. In the following, few selected examples of both stabilization and rigidification of proteins by genetic methods will be briefly mentioned. The stabilizing role of protein-bond calcium ions has been extensively documented with both mesophilic and thermophilic proteins (Roche & Voordouw, 1978; Fontana, 1991; Tainer et al., 1992; Nayal & Di Cera, 1994, and ref.es cited therein). Calcium binding is expected to induce and stabilize protein structure and consequently also to rigidify a protein molecule at the site of the ion binding, since Ca2+ can bind up to nine ligands. Themolysin binds four Ca2§ ions, while the thermolabile mesophilic protease from B. subtilis only two ions, despite the high sequence similarity between the two proteins (Roche & Voordouw, 1978). Genetic methods have been used by Toma et al. (1991) to replace a surface loop (from residue 188 to residue 194) of B. subtilis neutral protease with a 10-residue segment which, in the homologous polypeptide chain of thermolysin, binds Ca-4. The mutant protease was shown to bind an additional calcium ion and, moreover, to display enhanced stability in respect to the wild-type species. Engineering of calcium binding sites into proteins has proved to be aviable procedure for enhancing stability/rigidity of proteins (see Fontana, 1991; Tainer et al., 1992, for ref.es). A comparative analysis of amino acid sequences and three-dimensional structures of homologous proteins from mesophilic and thermophilic sources indicated that Gly--->Ala is the most frequent "cold to hot" amino acid exchange often occurring at helical segments (Argos et al., 1979; Menrndez-Arias & Argos, 1989; Vogt & Argos, 1997; Vogt et al., 1997). In the context of present discussion of rigidity of thermophilic enzymes, it is of interest to observe that Gly is the most flexible amino acid residue among the protein amino acid residues (Yan& Sun, 1997), while Ala is the best helixinducer (O'Neil & De Grado, 1990; Chakrabartty et al., 1991). Thus, the Gly--->Ala exchange appears to be a strategy followed by nature in engineering protein stability, as well as rigidity. The same amino acid replacement was proposed and utilized by Matthews et al. (1987) for enhancing the stability of T4 lysozyme, considering that the Gly~Xaa exchange is expected to indirectly stabilize the folded protein by decreasing the chain entropy of the unfolded state. This strategy of protein stabilization has been successfully employed by Margarit et al. (1992) for enhancing the thermostability of the mesophilic neutral protease from B. subtilis. Single and double Gly~Ala mutants of this
287 protease, which is homologous to thermolysin, were more stable than the wild-type protein. Moreover, it was shown that mutational effects were cumulative (Wells, 1990). The most rigid amino acid residue in proteins clearly is Pro, as a result of the steric constrains of the pyrolidine ring of this residue. Accepting the view that there is a correlation between rigidity and stability, one would predict that thermophilic enzymes should possess enhanced content of Pro residues. Indeed, Suzuki et al. (1987) has proposed the "proline rule" for thermophilic enzymes, in that they contain an enhanced number of Pro residues in respect to their mesophilic counterparts. The introduction of Pro residues into a protein was therefore utilized as a strategy for stabilizing proteins. Mutants of oligo-l,6-glucosidase from B. c e r e u s with multiple Pro exchanges (up to nine residues) were thus produced and shown to be significantly and cumulatively stabilized against thermal denaturation (Watanabe et al., 1994). The same Xaa~Pro replacement for enhancing protein stability was also proposed by Matthews et al. (1987) on the basis of entropic considerations of protein stability (see above). Since the Pro residue is a helix-breaker (O'Neil & De Grado, 1990), the Xaa~Pro replacement should occur at the level of loops in the protein molecule. If the same exchange is placed in the middle of a helical segment, the protein becomes less thermostable (Nakamura et al., 1997). The concept of protein stabilization by rigidification has been recently applied by De Filippis et al. (1998) introducing by a semi-synthetic procedure cx-amino-isobutyric acid (Aib, or cx-methylalanine) at selected positions in a folded 62-residue fragment of thermolysin (C-terminal subdomain 255-316). The geminal methyl groups of Aib severely reduce the conformational space of this amino acid residue and thus restrict the peptide backbone torsion angles to those characteristic of the helical configuration. Consequently, the Xaa---~Aib exchange is expected to rigidify and stabilize a protein in analogy to the Gly---~Ala or Xaa---~Pro exchange (Matthews et al., 1987). Indeed, a double Ala~Aib replacement in thermolysin fragment 255-316 enhanced the melting temperature from 63.5 to 71.5~ (ATm= 8~ (De Filippis et al., 1998). 7. DISCUSSION The rigidity of the stable enzymes that can be isolated from thermophilic and extremophilic microorganisms is documented by a variety of experimental findings. The major effect of this protein rigidity is in reducing the catalytic efficiency of thermophilic enzymes in respect to that of their mesophilic counterparts and thus thermophilic enzymes are usually poor catalysts at room temperature. It is proposed that a rigid protein structure does not allow correct proximity and positioning of the key residues required for an efficient catalysis, which requires instead a somewhat flexible protein architecture to bind and properly accomodate the substrate at the active site, to perform the catalytic evem and to release the product of the enzymatic reaction (Alber et al., 1983; Fersht, 1985). Considering that extremophiles appeared earlier during biological evolution (Stetter et al., 1990; Stetter, 1993), it can be proposed that the more flexible (and allosteric) mesophilic enzymes derived from thermophilic ones and, since at
288 ambient temperature catalysis is reduced, evolution created more efficient biocatalysts by improving their plasticity (for an alternative view see Danson et al., 1996). Having proposed that thermophilic enzymes are more rigid molecules, one should expect that psychrophilic enzymes should be even more flexible than the mesophilic ones. Indeed, the thermolabile enzymes isolated from psychrophilic microorganisms have been shown to possess improved catalytic activity in respect to that of their mesophilic or thermophilic counterparts. The molecular basis for this effect has been analyzed utilizing the structural models of some psychrophilic enzymes produced by homology modelling (Feller et al., 1996; Narinx et al., 1997) or by direct determination of their X-ray structure (Aghajari et al., 1996; Feller et al., 1996). In general, it has been observed that psychrophilic enzymes possess enhanced overall flexibility in respect to their mesophilic or thermophilic enzymes, as judged from physicochemical data (Feller & Gerday, 1997). It has been proposed that the structural factors favouring the conformational flexibility of psychrophilic enzymes likely reside in a reduced number of salt bridges, shorter chain loops, less proline residues in loops, low hydrophobicity of protein core(s) and weaker binding of calcium ion(s) (Feller et al., 1996; Narinx et al., 1997). In general, it can be stated that the results of structural, functional and stability studies on psychrophilic enzymes parallel those conducted on thermophilic and extremophilic enzymes, since the results of both studies indicate the same factors and interactions as modulators of protein stability/rigidity/flexibility (see Feller & Gerday, 1997; Gerday et al., 1997, for a review). As an example, a higher or lower content of Pro residues in loops appears to correlate with enhanced or reduced thermostability in thermophilic and psychrophilic enzymes, respectively (Feller & Gerday, 1997). In conclusion protein flexibility appears to be a main reason for the enhanced activity of psychrophilic enzymes, while protein rigidity explains why thermophilic enzymes are poor catalysts at room temperature. Nevertheless, enzyme catalysis is a complex process involving a proper stereochemistry of the active site, specific binding of the substrate, participation of functional (acid or basic) groups characterized by specific pK values, a microenvironment at the active site of defmed dielectric constant, etc. Since proteins must move in order to function, an efficient active site requires some mobility, but it is clear that the molecular mechanism of enzyme catalysis is the result of a subtle balance between a variety of energetic, structural and dynamic effects and, consequently, perhaps it is not appropriate to overemphasise the role of protein mobility in catalysis. We are inclined to consider therefore protein rigidity of thermophilic enzymes an often made observation, but we are aware that exceptions can exist. For example, a laboratory evolution technique has been devised by using an engineered E. coli containing the LeuB gene coding for the isopropylmalate dehydrogenase from the extreme thermophile Thermus thermophilus (Topt 80~ Upon prolonged incubation at moderate/ambient temperature, E. coli mutants able to grow rapidly were obtained, producing isopropylmalate dehydrogenase mutants which were shown to retain the unusual thermostability of the wild-type enzyme from Th. thermophilus, while displaying high catalytic activity even at ambient temperature (Suzuki et al., 1997). Since it is possible
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
295
I m p r o v e m e n t o f thermal stability o f a diagnostic e n z y m e , Streptomyces cholesterol oxidase, b y r a n d o m and site-directed m u t a g e n e s e s and a structural interpretation Y. Murooka a, Y. Nishiya b, M. Toyamaa , M. Aoike a, M. Yamashita ~, and N. Hirayama c aDepartment of Biotechnology, Graduate School of Engineering, Osaka University, Yamadaoka, Suita, Osaka 565-0871, bTsuruga Institute ofBiotechnology, Toyobo Co. Ltd., Toyo-cho, Tsuruga, Fukui 914, r of Biotechnology, Faculty of Engineering, Tokai University, Nishino, Numazu, Shizuoka 410-03, Japan
1. INTRODUCTION Cholesterol oxidase (313-hydroxysteroid oxidase, EC 1.1.3.6) is a FAD-dependent bifunctional enzyme that catalyzes both the oxidation of cholesterol (5-cholesten-3[3-ol) to the intermediate 5-cholesten-3-one with the reduction of molecular oxygen to hydrogen peroxide, and the isomerization of steroid with a trans A:B ring junction to reduce 4cholesten-3-one. This enzyme is industrially important, and is commonly used for the enzymatic transformation of cholesterol. It is also useful for the clinical determination of serum cholesterol by coupling with a related enzyme for the assessment of arteriosclerosis and other lipid disorders and of the risk of thrombosis. Another use of the enzyme is in the microanalysis of steroids in food specimens. We previously cloned (1) and sequenced (2) the gene for cholesterol oxidase (choA) from Streptomyces sp. SA-COO. The secretory overproduction of Streptomyces cholesterol oxidase (ChoAs) in a Streptomyces host-vector system was demonstrated (3). Recently, the expression of the choA gene from Streptomyces in Escherichia coli by genetic modification was also reported (4). The crystal structure of cholesterol oxidase (ChoAB) from Brevibacterium sterolicum was determined and refined at 1.8A resolution (5). Li et al. reported the structure of a complex of cholesterol oxidase with the steroid substrate dehydroisoandrosterone, refined at 1.8A resolution (6). The steroid is buried within the protein in an internal cavity which, in the free enzyme crystal structure, is occupied by a lattice of water molecules. The hydroxyl group of the steroid substrate is hydrogen-bonded to both the flavin ring system of the FAD cofactor and a bound water molecule. The overall topology of this domain is very similar to that of other FAD-binding proteins. The amino acid sequence of ChoAs is homologous with that of ChoAB (59.2% identity). In particular, residues composing the steroid- and the FAD-binding sites are highly conserved between ChoAs and ChoAB. The three-dimensional structure of ChoAs modeled on the basis of its homology to ChoAB provides a reasonable point for analyzing the structure-activity relationships of thermostable mutants of ChoAs created by random mutagenesis. Here we report on successful enhancement of the thermal stability of ChoA by means of in vitro random mutagenesis and by site-directed mutagenesis techniques. The mutational effects are discussed using a structural model of ChoAs based on the tertiary structure of
296 ChoAs. We also describe other effects of the amino acid substitutions, which provide information for further improvements in the functionality of enzyme. 2. MATERIALS AND M E T H O D S The host strain used was E. coli JMI09. Plasmids pCOll7 coding for C h o A (4) and pBluescript-KS were used for additional plasmid construction. Plasmid pCOll0 (7), prepared for random mutagenesis of the choA gene, contains a unique SphI site on the choA gene. Recombinant strainswere grown in Terrificbroth or on L agar. The antibiotic used was ampicillin(50 ttg/ml). The transformation of E. coli, isolation of the plasmid, cleavage of the D N A with restriction enzymes, polymerization with Klenow fragment and ligation with T4 D N A ligase were carried out according to the common method. The nucleotides were sequenced on an A L F D N A sequencer. Random mutagenesis of the choA gene was carried out using P C R techniques. P C R was done under standard conditions with Tth D N A polymerase (Toyobo Co., Osaka, Japan), except that dTTP or dCTP was one fifth the volume of other deoxyribonucleotidc phosphates. The amplified fragments and pCOl10 were digested with SphI and ApaI, respectively. The insert fragments and vector were separated by agarosc gel electrophoresis, purified and ligated. These constructs were used to transform E. coli JM109 and transformants were plated onto indicatorplates. Cholesterol oxidase-producing colonies selected were grown on L agar at 30~ and heat treated by incubation at 50~ for 16 h. After the heat treatment, positive clones having enzyme activitywere selected by the filter-paper method (1) based on a colorimetric assay of cholesterol with cholesterol oxidase and peroxidase (8). These isolates were thermostablc enzyme-producing clones, because E. coli JM109 (pCOl l0) producing the wild-type C h o A showed no activity after the heat treatment. Site-directed mutagenesis was performed with a Transformer kit (Clontech Laboratories, Palo Alto, CA) as described in the manufacturer's instructions. Successful mutations were verified by DNA sequencing. The mutagenesis oligonucleotides used were synthesized with a Model 381A DNA synthesizer (Applied Biosystems, Foster City, CA). Each recombinant strain was grown to the stationary phase at 30~ in Terrific broth with agitation at 180 r.p.m. Cells were harvested by centrifugation and crude extract was prepared by vortex mixing the cells with glass beads. Ammonium sulfate was added to the cell-free extract and the precipitate collected by centrifugation, dissolved in 50 mM phosphate buffer (pH 7.0) and dialysed against the same buffer. The enzyme was purified from the dialysate by DEAE-Sepharose column chromatography followed by gel filtration to homogeneity. Cholesterol oxidase activity was measured by the method of Allain et al. (8). Enzyme solutions (0.5-1.0 U/ml) were prepared by dilution with 20 mM potassium phosphate (pH 7.0) containing 0.2% bovine serum albumin. The appearance of quinoneimine dye formed by coupling with 4-aminoantipyrine, phenol and peroxidase was measured at 500 nm by spectrophotometry. One unit of activity was defined as the formation of 1 M o f hydrogen peroxide (0.5 ttM of quinoneimine dye) per minute at 37~ and pH 7.0.
297 3. RESULTS 3.1. Construction of ChoA structural model Sequence alignment was carried out using the Needleman-Wunsch algorithm (9). The ChoAs sequence showed a 59.2% homology with ChoAB. A complex between ChoAB and dehydroisoandrosterone, an inhibitor of cholesterol oxidase, determined by X-ray crystallography (6), provided a basis for three-dimensional structure modeling of ChoA (Figure 1). The ChoA model was constructed using the QUANTA software package (QUANTA 4.0; Molecular Simulations, Burlington, MA). The ChoAB coordinates were obtained from the Brookhaven Protein Databank (10). The initial model was refined by energy minimization using the steepest descent method followed by the conjugate gradient method (11). The minimization calculations were conducted using the CHARMm module of QUANTA. All calculations were performed on an Indy workstation (Silicon Graphics, Palo Alto, CA). 3.2. Random mutagenesis to isolate thermostable mutant cholesterol oxidases The thermostable cholesterol oxidase was isolated from recombinant cells carrying a randomly mutated gene. The SphI-ApoI fragment of pCO110 encoding a part of ChoA (from position 97 to position 355 in the precursor sequence composed of 546 amino acid residues) was used as the target DNA. Since the structural information of ChoAB, indicated that this region does not contain the major parts of the FAD-binding domain and the active site cavity (5), the enzyme activity of ChoA would be conserved by random mutagenesis of this region. Approximately 90% of recombinant cells carrying a randomly mutated gene showed cholesterol oxidase activity. After incubation at 50~ for 16 h, the cholesterol oxidase activities of four clones among 5000 colonies were detected on the indicator plates, although no activity was detected in the wild-type-producing strain, E. coli JM109 (pCOll0) under the same condition. Recombinant plasmids were extracted from the four clones and the DNA sequences of the inserted fragments were determined. A single base change, resulting in the replacement of one amino acid residue, was found in each choA gene. All of the amino acid substitutions were located within a limited region (SerlO3 to Thr, Va1121 to Ala, Arg135 to His and Vail45 to Glu). The four ChoA mutants were designated S103T, V121A, R135I-I and V 145E, respectively. 3.3. Characterizations of wild-type and mutant cholesterol oxidases The four recombinant strains producing the mutant ChoA were cultured and these cholesterol oxidases were purified to homogeneity. All the purified enzymes and the wildtype enzyme migrated as a single protein band in SDS-PAGE analysis; the molecular mass of each enzyme was determined to be-55 kDa. The specific activities ofS103T, V121A, R135H and V145E were found to be 86.6, 72.0, 68.9 and 63.5 U/mg, respectively, while that of the wild type was 67.7 U/mg (with 1.0 mM cholesterol as the substrate, at 37~ and pH 7.0). The mutant enzymes thus had specific activities similar to or slightly higher than that of the wild-type enzyme. The thermal stabilities of these mutant enzymes were compared with that of the wild-type enzyme. All the mutant enzymes were more thermally stable than the wild type at 50, 55 and 60~ (Figure 2). The half-lives of the wild-type, S103T, V121A, R135H and V145E were calculated to be 7.8, 11.3, 12.2, l l.0 and 24.1 min, respectively. The increased thermal stability of V145E was significant.
298
100 V145E
A
v
>,10 >
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..=,
IS103T R135H
ro~
c r
1
,Wild type
E
~D tr
"10 Figure 1. Representation of the overall folding of Stm/~omyces cholesterol oxidase that is constructed by hon~logy modeling. The FAD molecule (red balls) and de~droisoatxtm~ t m e (gray balls) are indicated. Typical mutation sites are also indicated.
"
I~O
40
60
Incubation time at 60"C (min) Figure 2. Effect of temperature on the stability of wild-type and mutant cholesterol oxidases.
The effects of pH on the stability and activity of the mutant and wild-type cholesterol oxidases were also compared. The stabilities of their pH were similar to that of the wildtype enzyme. However, the pH profile of VI45E differed from that of the wild-type and other mutant enzymes. The optimum pH of Vl45E was shifted to alkali and encompassed a broad range between acid and alkali. This significant alteration in pH in V145E may be caused by introduction of the negatively charged amino acid, glutamate. The Km values for cholesterol were calculated for the purified wild-type and mutant enzymes. The Km value for $103T, R135H and V145E, 13, 30 and 11 ~M, respectively, were similar to that for the wild-type enzyme, which was calculated to be 13 ~M. It was difficult to determine a K= value for V121A because the substrate-velocity curve was different from the Michaelis-Menten type. 3.4. Site-directed mutagenesis of V145 and other amino acid residues Since ChoAa was more thermostable than the ChoAs, we searched the differences of amino acid residues between ChoAs and ChoAa. We predicted some amino acid residues that might have effect on the stability of the enzyme and thus substituted V145, Q286, L119, P357 and $379 by other amino acid residues (Figure I). These mutant enzymes were also purified to homogeneity and characterized. The VI45D and VI45E mutants shiRed the optimum temperatures to 67~ but not other V145Q, Q286R, LII9A, LII9F, P357N and $379T mutant enzymes (Figure 3). The thermal stabilities of these mutant enzymes were compared with that of the wild-type enzyme (Figure 4). VI45E and VI45D were markedly thermostable. The V145 of ChDAs is corresponding to QI08 of ChDAB (Figure
299 120
~2o
100
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,~v* 8 0 -
o~~
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~(J6 0 \
Wild type
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~
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,
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.
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l
60
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80
Wild type L119A L119F P357N S379T l
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!
40
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50
!
60
i
70
Temperature (~
Figure 3. Optimum temperatures of wild-type and mutant cholesterol oxidases.
1201
120
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"~
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~
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~ -- ~"" "
-- i-
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,~
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a~ rr
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~
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"~
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- Q286R
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---On Wild type
----D--- L119A 2 0 ~ -- A--. Ll19F ----e--- P357N --L. S379T
o-t
70
30
40
50
60
70
Temperature (~
Figure 4. Thermal stabilities of wild-type and mutant cholesterol oxidases. The enzyme in 10raM ILP buffer (pH7.5) was incubated at various temperatures for 15 min.
!
80
300 5). The VI45Q mutation resulted in slightly increasing the optimum temperature and slightly stable than the wild-type. Q286R, L119F and $379T mutants were more thermally unstable than the wild-type. 3.5. Construction of multiple mutant enzymes A combination of mutations sometimes affecting protein stability shows additivity. Thus, we tried to construct multiple mutant enzymes by site-directed mutagenesis. The VI45E mutation was chosen as the base because of its higher level of thermal stability. As close residues easily influence one another, the S103T, V121A and R135H mutations were added in that order. Two-, three- and four-point mutant enzymes were designated M2 (S 103T and V145E), M3 (S103T, V121A and V145E) and M4 (SI03T, VI21A, RI35H and V145E), respectively. Each multiple mutant cholesterol oxidase was purified to homogeneity. These mutant enzymes were the same M.W. as that of the wild-type as determined by SDS-PAGE analysis. The specific activities of M2, M3 and M4 were found to be 39.9, 47.8 and 26.7 U/mg, respectively. The stabilities of M2 and M3 were elevated compared with that of V145E (Figure 6). The half-lives of M2 and M3 were 32.4 and 52.2 min, respectively, showing that the mutational effects of amino acid positions 103, 121 and 145 are additive. The stability of M4, however, decreased in comparison with those of V145E, M2 and M3.
100
S103T, V121A, V145 E (M3)
~.
'S103T, V145E (M2) VI45E (MI)
~> 10
:~ fo r U~ r
S103T,V121A, R135H,V145E (M4)
E ID al
-
9
20
9
40
-
9
60
Incubation time at C~'C (rain) Figtae 5. View of the residues arot~ V145 o f ~ p t o m ) r ~ C~A(A) arid
Figure 6. Effect of temperature on the stability of multiple mutant cholesterol oxidases.
O 108 ofBrev/bacterqwn C~A. Moreover, M4 exhibited maximum activity at 50~ whereas the optimum temperature of other enzymes was 60~ Thus, the R135H mutation has a negative influence on the other mutations. The optimum pH of the multiple mutant enzymes changed to encompass a broad range. The Km values of M3 and M4 for cholesterol could not be estimated because the substrate-velocity curves were similar to that of mutant VI21A. The Km value of M2 was calculated to be 21 M . The multiple mutant enzymes thus inherited the features of the V145E and/or V121A mutants.
301 4. DISCUSSION
We succeeded in creating thermostable cholesterol oxidases by random and site-directed mutageneses. The three-dimensional structure of ChoA modelled on the basis of its homology to ChoAB helps to analyze the structure-stability relationships of the mutants. The structural model also helps us to deepen our understanding of the reaction mechanism of cholesterol oxidase. Replacement of the V145 by E or D could introduce not only a hydrogen bond between E145 or D145 and D134, but also a salt bridge between E145 or D145 and R147 (Figure 5). These newly formed attractive interactions could contribute to the stabilization of the conformation and increase the thermal stabilities of V145E and V145D. The interactions of E145 with D134 and R147 would certainly influence the deprotonation and protonation of D134 and R147. The drastic change in the optimum pH ofV145E is considered to be due to these interactions. The two residues corresponding to positions 135 and 145 of ChoA lie on each of two antiparallel 13-strands and are located near each other. Replacement of these residues by H and E, respectively, in M4 will cause a very short non-bonded contact between an atom at the ~osition of H and C• of E. The interaction would destructively influence the molecular conformation there and reduce the thermal stability. M2, containing two point mutations, was finally selected as the commercial type of ChoA. Although three point mutations, M3 was the most thermostable mutant, the substrate-velocity curve was different from that of Michaelis-Menten type on account of the V121A mutation. Therefore, M3 was unsuitable for application of enzymatic assay. Arising from the effect of the V145E mutation, M2 exhibits a broad range of optimum pH from acid to alkali. This makes the variant suitable for clinical application, because various buffers and pHs are encountered depending on the assay reagent components. Development of M2 for commercial production is now in progress. Recently, we have succeeded in creating new enzymes that catalyze either the oxidation of cholesterol or the isomerization of 5-cholesten-3-one, as well as in enzyme improvement with respect to either or both reaction rates by the substitution of some amino acids. These findings will provide several ideas for the design of more powerful enzymes.
REFERENCES
1. 2. 3. 4. 5. 6. 7. 8.
Y. Murooka, T. Ishizaki, O. Nimi, and N. Maekawa, Appl. Environ. Microbiol., 52 (1986) 1382-1385. T. Ishizaki, N. Hirayama, H. Shinkawa, O. Nimi, and Y. Murooka, J. Bacteriol., 171 (1989) 596-601. I.'Molnar, K.-P. Choi, N. Hayashi, and Y. Murooka, J. Ferment. Bioeng., 72 (1991) 368-372. N. Nomura, K.-P. Choi, M. Yamashita, H. Yamamoto, and Y. Murooka, J. Ferment. Bioeng., 79 (1995) 410-416. A. Vrielink, L.F. Lloyd, and D.M. Blow, J. Mol. Biol., 219 (1991) 533-554. J. Li, A. Vrielink, P. Brick, and D.M. Blow, Biochemistry, 32 (1993) 11507-115151 Y. Nishiya, N. Harada, S.-I. Teshima, M. Yamashita, I. Fujii, N. Hirayama, and Y. Murooka, Protein Eng., 10 (1997) 231-235. C.C. Allain, L.S. Poon, C.G.S. Chan, W. Richmond, and P.C. Fu, Clin. Chem., 20 (1974) 470-475.
302 9. S.B. Needleman and C.D. Wunseh, J. Mol. Biol., 48 (1970) 443-453. 10. F.C. Bemstein, T.F. Koetzle, G.J.B. Williams, E.F. Meyer, M.D. Brice, J.R. Rodgers, O. Kennard, T. Shimanouchi, and Y. Tasumi, J. Mol. Biol., 112 (1977) 535-542. 11. I. Fujii, N. Hirayama, M. Yamashita, and Y. Murooka, Abstract Book of Annual Meeting of Nippon Seibutsu Kougaku (the Society for Fermentation and Bioengineering, Japan), (1994) 106.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
303
Protein engineering for thermostabilisation of proteins: some theoretical rules and application to a [3-glucanase E. Querol a, J. Pons a, J. Cedano a, M. Vallmitjana a, F. Garcfaa, C. Bonet b, J. P6rez-Pons a A. Planas c & A. Mozo-Villarfas b alnstitut de Biologia Fonamental and Departament de Bioqufmica i Biologia Molecular. Universitat Autbnoma de Barcelona. Bellaterra. 08193 Barcelona. Spain. bDepartament de Ci~ncies M~diques B~siques, Facultat de Medicina, Universitat de Lleida. E25198 Lleida, Spain. CLaboratori de Bioqufmica. Institut Qufmic de Sarri~. Universitat Ramon Llull. 08017 Barcelona. Spain.
Protein thermostability has been investigated by two approaches. (A) Computational. To study the relationship between thermostability and conformational characteristics of proteins, 195 single amino acid residue replacements have been analysed for several protein conformational characteristics. From the analyses, some general rules arise which suggest where amino acid substitutions can be made to enhance protein thermostability. (B) Experimental. Glucohydrolases are biotechnologically important enzymes. We are analysing by site directed mutagenesis the structure/function relationship of two bacterial glucohydrolases, a 1,3-1,4-13-glucanase and a 13-glucosidase. We have determined the key residues for catalysis and substrate binding, and redesigned the stability and specificity of the glucanase. A glucanase thermorresistant mutant (N57A) has been obtained. 1. INTRODUCTION An important goal of protein engineering for biocatalysis is the enhancement of protein thermostability. The use of thermostable enzymes in industrial applications offers the benefits of increased rates of reaction, higher substrate solubility, decreased media viscosity, longer enzyme shelf lives than at normal storage temperatures and lowered risk of microbial contamination when reactions are carried out at high temperatures.
However, although a
number of successful examples of stabilisation of proteins exist, the mechanism of their thermostabilisation is far from being understood (see [ 1] for a review). Up to the present moment the successful examples of stabilisation of proteins have been achieved by methods based not far from intuition or trial and error. Two general approaches have been followed to
304 analyse the stability of proteins. First, the comparison between homologous proteins from thermophiles and mesophiles has provided some insight into the reasons why related proteins performing the same functions could have very different stability. Second, the analysis by protein engineering of the contribution of the different interactions that take place in a protein has resulted in some general rules about possible ways to increase the stability of a protein [2 11]. The first approach has the limitation that it is often difficult to identify the important determinants involved in a specific case since, in general their sequences have diverged significantly. It is important to examine proteins that are at the beginning of divergence, when their structures are still almost identical and the effect of single differences can be easily analysed. Following the second approach, protein engineering, several strategies have been proposed, for example: (a) reducing the difference in entropy between folded and unfolded proteins, which in practice means reducing the number of possible conformations in the unfolded state, (b) stabilising the dipoles of m-helices, (c) increasing the number of hydrophobic interactions and packing ratio in the interior core and, (d) reducing the area of water-accessible hydrophobic surface. Nevertheless, simple patterns that characterise stabilising interactions and quantitatively predict the effects of amino acid substitutions have eluded identification. An additional problem is the fact that in interpreting the stabilisation in terms of specified local interactions, one has to consider that the free energy of stabilisation of globular proteins in solution represents a marginal difference of large numbers. Thus, important contributions coming from small rearrangements of side chains or structural elements may escape detection. The difference in AAG between mesophilic and thermophilic enzymes is about 5-7 kcal.mo1-1, which can be accomplished by a few hydrogen bonds or by two salt bridges inside the protein globule.
This marginal energy for stabilisation of proteins is clearly a result of natural
selection. There appears to be a selective pressure for the marginal stability, probably to facilitate such processes as polypeptide folding and the flexibility required by the "native" conformation in the protein function. On the other hand, there is no selective pressure to make proteins stable at temperatures above those which they encounter in vivo. In order to gain insight into a variety of structural and physico-chemical characteristics leading to thermostabilisation several properties were analysed, suggesting some general rules for stabilization. In addition, they have been applied to the redesign of glucohydrolases. Glucohydrolases are biotechnologically important enzymes. We are analysing, by protein engineering, the structure/function relationship and determining the key residues for catalysis and substrate binding, and redesigning the stability and specificity of two enzymes, 1,3-1,4-13D-glucan 4-glucanohydrolase (EC 3.2.1.73) and a ~l-glucosidase (EC 3.2.1.21). These enzymes are utilized in the food industry, in brewing, feedstuffs, for the enzymatic synthesis of carbohydrates or glycoconjugates favoring their transglycosylating activity, etc. Among other
305 site-directed mutants with improved catalytic efficiency, etc., we have designed 13-glucanase thermorresistant mutants. We have previously determined the key catalytic residues by protein engineering and the 3D structure by crystallography [ 12 - 15]. 2. MATERIALS AND METHODS 2.1. Data from thermostable and mesostable isoproteins The wild type and thermostable mutant proteins used in this database are: o~-amylase; o~lantitrypsin; calbindin; carboxyl esterase; chloramphenicol acetyltransferase; cytochrome C; ferredoxin; [~-galactosidase; glucose dehydrogenase; glyceraldehyde phosphate dehydrogenase; 3-isopropylmalate dehydrogenase; kanamycin nucleotidyltransferase; lactate dehydrogenase; T4, hen egg-white, bird and human lysozymes; luciferase; malate dehydrogenase; neutral protease; oligo-l,6-glucosidase; protein HU; ~ cro and arc repressors; ribonucleases A, T1 and barnase; RNA polymerase subunit; subtilisins; superoxide dismutase; triosephosphate dehydrogenase; tryptophan synthase; tyrosinase; xylanase; xylose isomerase. The bibliography used to obtain this data is reported in a previous article [ 10]. Most of the thermostable forms have been obtained by site-directed mutagenesis, few of them emerge from the analysis of native isoproteins, in which the mesostable and thermostable forms differ only in one amino acid residue. From 195 single amino acid residue replacements -164 different positions on protein sequences- reported elsewhere [ 10] several protein conformational characteristics have been analysed. These are: type of residue replacement; conservative versus non conservative substitution; replacement being in a homologous stretch of amino-acid residues; change in hydrogen bonds, van der Waals and secondary structure propensities; solvent accessible versus inaccessible replacement; type of secondary structure involved in the substitution; the physico-chemical characteristic to which the thermostability enhancement can be attributable and, finally, the relationship of the replacement site to the folding intermediates of the protein, when known. Details of the computer analysis are described elsewhere [ 10]. 2.2. Experimental Bacterial strains, chemicals, PCR site directed mutagenesis, protein purification, enzyme kinetic assays, equilibrium urea denaturation, enzyme inactivation and thermotolerance (ts0) measurements, have been performed as previously reported [ 13, 15 - 17]. 3. RESULTS AND DISCUSSION. 3.1. Theoretical results. From the 195 substitutions (164 different positions on protein sequences), 121 represent a
306 different type of residue replacement. Since the whole space of directional changes has 380 substitutions (20 x 19), a quite large set of the potential substitution space has been explored by Nature or by protein engineers. As could be expected (see Table I), most replacements are conservative according to the Dayhoff matrix of replacements [ 18]. Table I. Some propensities related to thermostable replacements Secondary structure Potentiality for Conservative (Dayhoff) propensity hydrogen bonding Conserved Non conserved Maintainor Decrease Maintain or Decrease increase increase 146/195 49/195 162/195 33/195 121/195 74/195 It was also found that most replacements lead to maintain or increase the secondary structure propensity, analysed according to a classical scale of secondary structure propensities [19]. Most of the replacements lead to a residue with the same or enhanced potential for hydrogen bonding lying on its side chain, independently of their sequential and structural context. This is in agreement with a recently reported analysis [ 11]. The type of secondary structures involved in the replacements are shown in Table II. As can be seen, most of them lie in regular secondary structure positions either in a-helices or in [5-strand. Table II. Secondary structure involved Total a-helix 6 13-strand turn loop coil
3 2 2 2
On the N-cap 16
.....
On the C-cap 6
6 nd
3 nd
nd nd
nd nd
nd = No data available
If the above results are explainable in terms of conservation of protein structure and function, the results related to the rate of external versus internal replacements are more difficult to explain. Of 164 mutations, 121 were done in solvent accessible regions and 43 were on buried regions. This could be related to the fact that mutations within the core of a protein are likely to affect more than one energy term. Therefore evolution could choose those mutations being conformationally less expensive, which correspond to residues at the solvent exposed area. The putative amphipathicity of the regular secondary structures in which the mutants lie has also been checked. From the 109 point mutations, 23 lie on different amphipathic a-helix or ~-strand secondary structures (19 on helices and 4 on strand). However, this result may also
307 reflect the biased selection done by researchers. We have checked whether the internal replacements would primarily correspond to internal residues from amphipathic structures. We have found the following: 15 external, 16 internal amphipathic-helix residues, and 4 external, 2 internal amphipathic-strand residues, results that do not suggest any special tendency. Another interesting result comes from the homology/similarity analyses of the amino acid sequence stretches in which the mutated residue lies (we do not refer to the conservation of the specific replaced residue, which has been analysed above). This has been done by the Clustal algorithm [20] on those proteins having a number of sequences to allow this analysis (in fact it was the whole set of proteins used in this work except lysozymes and repressors); 113 regions in total as seen in Table III: Table III. Replacement on conserved stretches of mult!ali~ned homologous sequences On regular secondary structure On loops, turns or coils Highly conserved region 46 36 Medium conserved region 6 10 Non conserved 7 8 ,
,
,
Most of the replacements match conserved stretches of residues, specially those lying on a regular secondary structure. It is a well known fact that regular secondary structure stretches are highly conserved in protein sequences, however to our surprise most substitutions lying on loops, turns and coil regions, correspond to conserved regions too. Thermostability is certainly related to protein structure and conserved external regions are important for structure indeed. It is usually assumed that replacements leading to enhanced protein thermostability correspond to rigid regions. We have checked it for those cases in which these data (B-factors) are either reported by the authors or can be obtained from the Protein Data Bank. We found 37 replacements in flexible regions, 38 in rigid regions (113 were found in regions on unknown rigidity), suggesting that a simple increase of rigidity does not necessarily lead to thermostablity enhancement. Finally, a survey of the physicochemical basis of the thermostability enhancement, when indicated by the authors, show that they are diverse. This list is biased by the fact that sitedirected mutants are chosen by researchers to accomplish, as far as possible, some physicochemical expectations. Table IV summarises these results. In a subset of replacements leading either to a thermostability enhancement of 5~ or higher, or to a doubling of the thermoresistence, we have found an outstanding improvement of secondary structure propensity with respect to the whole set of 195 substitutions: Most of these substitutions lie on conserved and solvent-accessible regions.
308 Table IV. Physical and chemical contributions to AG to which thermostability enhancement be attributed "Better hydrogen bonding Better hydrophobic internal packing Enhanced secondary structure propensity Helix dipole stabilization Argo's replacements Removal of residues sensitive to oxidation or deamidation Burying hydrophobic accessible area Improved electrostatic interactions Strengthening intersubunit association Decrease chain strain Salt bridge optimisation Better Van der Waals Better affinity for calcium Improved AH upon substitution Unspecified improved AG upon substitution Unknown or not described
can 18 16 12 10 10 10 7 6 6 5 4 3 2 1
25 60
From the above results we suggest the following general rules to enhance protein thermostability: - Look for conserved stretches of residues upon multiple alignment. - Make substitutions conservative according to the Dayhoff matrix. Maintain or enhance the secondary structure propensity upon substitution.
-
- Replace preferentially solvent accessible residues. - Replace residues on N- and C-cap of m-helices (PDB or predicted) by introducing, if possible, a negative charge at N-cap or positive at C-cap of the helix. Replace residues on the N-cap and C-cap of l~-strands introducing, if possible, a negative -
charge at N-cap or positive at C-cap of the strand. Substitute with residues which improve or increase the number of potential hydrogen bonding
-
or van der Waals contacts. Whenever possible consider thermal damage on specific amino acid residues: deamidation of
-
Asn and Gin (the sequential motive Gly-Asn is a well known deamidation target [21]); oxidation of methionines; splicing the peptidic bond besides aspartic residues and substitutions of free thiol cysteins [21 - 25]. And also consider some suggestions elsewhere reported, as for example: - Argos' replacements (Lys ~ Arg, Asp ~ Glu, Gly ~ Ala) [2]. -
Substitute with prolines in loops [26].
- Introduce an additional disulfide bridge [27-29]. -
Introduce an additional metal binding site [30].
- Introduce new glycosylation sites [31].
309 Obviously, the best strategy for increasing the stability of a specific protein by sitedirected mutagenesis would require a thorough knowledge of its three-dimensional structure and of the molecular mechanisms responsible for its heat inactivation. There is still a limited number of proteins that fulfill these requirements. Since the above rules only predict wide regions of the protein as targets for site directed mutagenesis, in order to design individual replacements we have checked three different computational methods which define protein pseudopotentials. We find that each performs well only for one protein.
3.2. Site directed mutagenesis of a bacterial [3-glucanase The selected stretch of residues to be mutated constitutes a major loop covering the active site cleft. It is a solvent exposed region, highly conserved on most of the Bacillus glucanases (Figure 1). In addition to enhance thermostability, we were interested on redesigning and improving the catalytic and specificity properties of the enzyme (for more information see [ 15]). The general strategy was an Alanine scanning mutagenesis of the whole region followed by saturation mutagenesis of selected residues.
B. l i c h e n i f o r m i s B. m a c e r a n s B. p o l y m y x a B. subt ilis B. a m y l o l i q u e f a c i e n s
51 64 DGYSNGNMFNCTWR DGYSNGGVFNCTWR DGYSNGQMFNCTWR DGYSNGNMFNCTWR DGYSNGDMFNCTWR
Figure 1. Alignment of the loop amino acid sequences from different Bacillus glucanases. The designed single point mutations were: D51A, G52A, Y53A, $54A, N55A, G56A, N57A, M58A, F59A, N60A, T62A, W63A, R64A, M58D, M58S, M58V, M58W, M58G, M58F, M58N, M58P, M58L, M58I, M58Q, M58R, M58E, M58Y, M8T. A second strategy was the introduction of an additional disulfide bridge embracing the N- and C-terminal tails of the protein structure, by means of the double mutant $34C+K242C. Thermotolerance was determined at 65~ as the incubation time required to irreversibly inactivate the enzyme to 50% of its initial activity (ts0). One mutant, N57A showed higher thermotolerance (70%) than the wt enzyme. It is well known than Asn residues are often involved in protein deamidation. This effect is more pronounced when Asn is next to a Gly residue in a highly solvent exposed region [21 ], which is the case in this glucanase. Therefore, we suggest than the thermotolerance is due to removing the Asn residue. Finally, the strategy of introducing an additional disulfide bridge by means of the double mutant $34C+K242C failed. We got, at lower expression levels than for wt and other mutants, a protein with the same thermoresistence than the wild type. Finally, it deserves to be mentioned that, in addition to enhance thermostability, we were interested on
310 redesigning and improving the catalytic and specificity properties of the enzyme (for additional information see [15]). ACKNOWLEDGMENTS
This research was supported by grants BIO97-0511-CO2 and IN94-0347 from the CICYT (Ministerio de Educaci6n y Ciencia, Spain) and by the Centre de Refer~ncia de R+D de Biotecnologfa de la Generalitat de Catalunya. REFERENCES.
1. Gupta, M. N. (ed.) (1993) Thermostability of enzymes. Springer-Verlag, Berlin. 2. Argos, P., Rossmann, M.G., Grau, U. M., Zuber, H., Frank, G. and Tratschin, J. D. (1979) Biochemistry 18,5698. 3. Imanaka, T., Shibazaki, M. and Takagi, M. (1986) Nature, 324, 695. 4. Querol, E. and Parrilla, A. (1987) Enzyme Microb.Technol., 9, 238. 5. Menendez-Arias, L. and Argos, P. (1989) J. Mol. Biol., 206, 397. 6. Mrabet, N. T., Van der Broeck, A., Van den Brande, I., Stanssens, P., Laroche, Y., 20. Higgins, D.G. and Sharp, P. M. (1989) Comp. Appl. Bioscien., 5, 151. 7. Fersht, A. R. and Serrano, L. (1993) Curr. Opin. Struct. Biol., 3, 75. 8. Nosoh, Y. and Sekiguchi, T. (1993) In Gupta, M. N. (ed.) Thermostability of enzymes. Springer-Verlag, Berlin, p. 182. 9. Serrano, L., Day, A. G. and Fersht, A. R. (1993) J. Mol. Biol., 233,305. 10. Querol E., P6rez-Pons J.A., Mozo-Villarfas A. (1996) Prot. Engineer., 9,265. 11. Vogt G., Woell S., Argos P. (1997) J. Mol. Biol., 269, 631. 12. Planas A., Juncosa M., Lloberas J., Querol E., FEBS Letters (1992), 308, 141. 13. Juncosa M., Pons J., Dot T., Querol E, Planas A. (1994), J. Biol. Chem. 269, 14530. 14. Hahn M., Pons J., Planas A., Querol E., Heinemann U. (1995), FEBS Letters 374,221. 15. Pons J., Querol E., Planas A. (1997) J. Biol. Chem. 272,13006. 16. Juncosa M, Pons J., Planas A., Querol E. (1994), BioTechniques 16, 820. 17. Pons J., Querol E., Planas A. (1995), Prot. Engineer. 8, 939. 18. Dayhoff, M. O. (ed.), (1978) Atlas of Protein Sequence and Structure. Natl. B iomed. Res. Found., Washington DC. Vol 5. Suppl. 3. 19. Privilege, P., Fasman, G. (1989) In Fasman, G. (ed.) Prediction of protein structure and the principles of protein conformation. Plenum Press, New York, 391. 21. Volkin D.B., Mach H., Middaugh C.R. (1995) Meth. Mol. Biol., 40, 35. 22. Ahern, T. J. and Klibanov, A.M. (1985) Science, 228, 1280. 23. Zale, S. E. and Klibanov, A.M. (1986) Biochemistry, 25, 5432. 24. Volkin, D. B. and Klibanov, A. M. (1987) J. Biol. Chem., 262, 2945. 25. Tomazic, S. J. and Klibanov, A. M. (1988) J. Biol. Chem., 263, 3086. 26. Matthews, B. W., Nicholson, H. and Becktel, W. J. (1987) Proc. Natl. Acad. Sci. USA, 84, 6663. 27. Perry, I. J. and Wetzel, R. (1984) Science, 226, 555. 28. Pantoliano, M. W., Ladner, R. C.,Bryan, P. N., Rollence, M. L., Wood, J. and Poulos, T. L. (1987) Biochemistry, 26, 2077. 29. Matsumura, M., Signor, G. and Matthews, B. W. (1989) Nature, 342, 291. 30. Toma, S., Campagnoli, S., Magarit, I., Gianna, R., Grandi, G., Bolognesi, M., DeFilippis, V. and Fontana A. (1991) Biochemistry, 30, 97. 31. Olsen, O. and Thomsen, K. K. (1991) J. Gen. Microbiol., 137, 579.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998Elsevier Science B.V. All rights reserved.
Mechanisms of hyperthermophilic
stabilization of the ~-glycosidase a r c h a e o n S ulfolobus solfataricus
311
from
the
M. Moracci a, M. Ciaramella a, L.-H. Pearl b, B. Cobucci Ponzano a and M. Rossia, c aInstitute of Protein Biochemistry and Enzymology, CNR, Via Marconi 10, 80125, Naples, Italy" bStructural Biochemistry Section, Department of Biochemistry and Molecular Biology, University College London, Gower Street, London, WCIE 6BT, United Kingdom CDepartment of Organic Chemistry and Biochemistry, University of Naples "Federico II", Via Mezzocannone 16, 80100, Naples, Italy
1. I N T R O D U C T I O N The interest arose in these years for enzymes and proteins from hyperthermophilic microrganisms comes both from the biotechnological advantages offered by stabilised biocatalysts or because the molecular basis of protein stability is a central problem in protein biochemistry far from being completely solved. In this regard, enzymes that are active and stable at temperatures up to 100~ have been chosen as model system for studying the adaptation of molecules to high temperatures. Thermostability in proteins is the result of the balance of two opposite factors: flexibility and rigidity (Jaenicke et al. 1996). Even few weak interactions, as five hydrogen bonds or salt bridges, can contribute significantly to stability. The comparison of the amino acid sequences of thermophilic and mesophilic
This work was supported by CNR Target Project Biotechnology and Bioinstrumentation, by CNR Progetto Finalizzato Ingegneria Genetica, by EC project: "Biotechnology of extremophiles" contract n. BIO2-CT93-0274and by EC FAIR project: "Enzymatic Lactose Valorization" contract n. 1048/96.
312 molecules provided indications on some preferences in terms of the amino acid composition of proteins from hyperthermophiles. However, these results were descriptive and could not explain in detail the molecular mechanisms of stabilization. More recently, the increasing number of 3D-structures of enzymes and proteins from hyperthermophiles (Day et al. 1992; Russell et al. 1994; Hennig et al. 1995; Kornd6fer et al. 1995; Yip et al 1995) allowed to shed some light on the determinants of structural stability. One of the thermophilic enzymes more extensively studied is the ~-glycosidase (EC 3.2.1.x) from the extreme thermoacidophilic archaeon Sulfolobus solfataricus (Ssl3-gly). This enzyme was purified to homogeneity, its gene was cloned and expressed in yeast, and the enzyme was fully characterised. It is a retaining glycosyl hydrolase with wide substrate specificity and remarkable exo-glucosidase activity against oligosaccharides, which are hydrolysed from the non-reducing end (for a review on this enzyme see Moracci et al. 1994). The amino acid sequence places the enzyme in glycosyl hydrolase family-1 along with archaeal, bacterial and eukaryal enzymes. More recently, the residues involved in catalysis were identified and the 3D-structure resolved at 2.6 ~ (Moracci et al. 1996; Aguilar et al. 1997). Like other enzymes from hyperthermophilic Archaea, Ss~-gly is extremely stable to heat, with an half-life of 48 hours at 85~ and displays optimal activity at temperatures higher than 85~ These features, together with the availability of the 3D-structure, make of SsI3-gly an attractive model system for the study of the structure-function relationship in proteins at high temperatures.
2. THE 3D-STRUCTURE OF SS~-GLY Ss[~-Gly has been crystallized in its native, tetrameric form using multiple isomorphous replacement, and refined at 2.6 ~ resolution. The enzyme shows the classic ([~o08 fold that has also been observed in the two mesophilic members of the glycosyl hydrolase family-1 crystallized so far: the cyanogenic [3-glucosidase from Trifolium repens (Barret et al. 1995)and the phospho-[3-glucosidase from
Lactococcus lactis (Weismann et al. 1995). The Ss[3-gly enzyme crystallizes as a tetramer with a dimer in the asymmetric unit; each monomer contacts two other monomers in the tetramer through extensive interactions involving the last 25 residues at the C-terminal. The intermolecular contacts involving the C-terminal strand appear to contribute significantly to the overall stability of the enzyme.
313
2.1 Factors producing thermostability Several theories, mainly based on the comparison of the amino acid sequences of thermophilic and mesophilic molecules, have been proposed in order to explain the molecular origins of the enhanced stability of enzymes from hyperthermophiles. When this analysis is applied to Ss[~-gly it is clear that most of these rules do not hold, since the enzyme shows a lower content of alanine and isoleucine and an higher content of tryptophan and asparagine than the mean of the mesophiles (Aguilar et al. 1997). In contrast, two structural features might contribute to thermal stabilization in Ss~-gly. This enzyme maintains a surprising high number of buried hydrophilic cavities than is generally observed, with one buried solvent molecule on 11.4 amino acid residues versus a 1:27 ratio in proteins in general. This feature has only been observed so far in Ss~-gly. The second one is the high number of ionic groups involved in ion-pairs. The Ss~-gly tetramer contains 524 charged groups including the (z-amino and the carboxyl groups; 58% of these are involved in ion-pair interactions and around 60% of them occur as part of multiple ion-pair networks with at least three charge centres. The networks tend to occur in non contigous positions covering the surface and spanning different domains and subunits. In this way, the networks could act as an electrostatic cross-link between folded structural elements. The abundance of ion-pairs arranged in large networks has been found in other enzymes from hyperthermophiles (Hennig et al. 1995; Yip et al 1995). Interestingly, arginines occur frequently in ion-pair networks for their ability to form multidentate interactions: this might explain the higher number of arginines found in enzymes from hyperthermophiles. These findings suggest that large networks of ion-pairs are important for hyperthermostability; however, although the stabilizing effect of salt bridges in proteins was proposed more that 20 years ago (Perutz and Raidt, 1975), how electrostatic interactions contribute to protein stability is still object of debate (Goldman 1995). Recent papers showed that ion-pairs do not affect the thermostability of several hyperthermophilic enzymes (Russell et al. 1994). However, Jaenicke's group has recently demonstrated the role played by a ionic network in the stabilization of the Dglyceraldehyde-3-phosphate dehydrogenase from Thermotoga maritima (Pappenberger et al. 1997). Hence, the importance of ionic networks in each hyperthermophilic enzymes have to be checked experimentally. In Ss[~-gly, site-directed mutagenesis was applied in order to test the stabilizing effect of the largest network which occurs at the tetrameric interface between the carboxy termini. This ionic network involves 16 ion-pairs interactions in which
314 the penultimate Arg488 of each monomer is hydrogen bonded to the side chain of Glu345 in the same monomer and salt bridged to the (x-carboxyl group of the Cterminal His489 of a non-crystallographic symmetry related monomer (Figure 1).
Figure 1. Cartoon of the Ss~gly tetrameric
interface.
For
clarity, only two, out of four, Arg488-His489
interactions,
are highlighted.
3. COMPARISON OF WILD-TYPE AND MUTANT SS[~-GLY The mutation Arg488Ala was introduced by site-directed mutagenesis. No significant differences in behavior during purification could be detected; the steady state kinetic constants and the profile of thermal activation of the m u t a n t were similar to those of the wild type, suggesting that the mutation caused only minor alterations of the native structure.
3.1 Inactivation kinetics
The kinetic of thermal inactivation of the wild type and Arg488Ala m u t a n t enzymes was followed by incubating the samples at 75~
and at 85~
increasing time spans, and measuring the residual activity at 65~ The wild type Ss[~-gly, incubated at 75~
for
(Figure 2A).
after an activation in the first 2.5 hrs,
maintained 80% of the initial activity after 5 h. The half-lives were 24 h and 2 hrs at 75~
and 85~
respectively. In contrast, the mutant Arg488Ala showed a
significant reduction of the thermal resistance with half-lives of 3.5 hrs and 10 min at 75~ and 85~
respectively. At 85~ no activation could be found for both
315 the enzymes. Interestingly, when both the wild type and the mutant enzymes were incubated in 8M urea at 20~ no inactivation was observed for more than 8 hrs (data not shown). The thermal inactivation at 75~ with sodium chloride 4M is shown in Figure 2B: the denaturation profile of the wilde type Ss~-gly is comparable with that
obtained at 85~ without salt, with an half-life of 2.5 hrs. In contrast, the half-life of the Arg488Ala mutant, at these conditions, was 50 min. A
B
120
120
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100 i ~
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~ ..=
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% 0000
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o
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.
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300
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time (rnin)
time (rain)
Figure 2. Thermal resistance of wild type (circle) and mutant (square) Ss~-gly. (A) 75~ (open symbols) and 85~ (closed symbols). (B) 75~ and 4M NaC1. These results show the importance of the Arg488 residue in the thermal stabilization and suggest that the inactivation of the mutant is caused by perturbation of the ionic network at the tetrameric interface. In fact, sodium chloride resulted more effective than urea on Ss~-gly, indicating the prevalence of ionic interactions in the stabilization of this enzyme. Furthermore, the faster salt-inactivation observed for wild type Ss~-gly, if compared with the mutant, can be explained assuming that the wild type, with an intact ion-pair network, is more
sensitive
than the Arg488Ala mutant
to the
increased
counterion
concentration.
4. CONCLUSIONS
The strong denaturating effect of the ionic strength has already been shown on hyperthermophilic enzymes indicating the importance of ion pairs in thermal
316 stabilization (Tomschy et al. 1994). Here, heat and salt-induced inactivation experiments on the wild type and mutant enzyme, give support to the observation derived from the 3D-structure, that a large ion pair network at the tetrameric interface stabilizes Ss[3-gly. Studies are in progress to identify the contribution to stability of additional residues of the network, and to analyze the structural effect of such mutations by kinetic and equilibrium denaturation.
REFERENCES
Aguilar C., Sanderson I., Moracci M., Ciaramella M., Nucci R., Rossi M. and Pearl L.-H., J. Mol. Biol., 271 (1997) 789 Barret, T., Suaresh, C.G., ToUey, S.P., Dodson, E.J., and Hughes, M.A., Structure, 3 (1995) 951 Day M.W., Hsu B.T., Joshua-Tor L., Park J.B., Zhou Z.H., Adams M.W.W. and Rees D.C., Protein Sci., 1 (1992) 1494 Goldman, A., Structure, 3 (1995) 1277 Hennig, M., Dairmont, B, Sterner, R., Kirschner, K., and Jansonius, J.N., Structure, 3 (1995) 1295 Jaenicke R. Schuring H., Beaucamp N. and Ostendorp R. Adv. Protein Chem., 48, (1996) 181 Kornd6fer, I., Steipe, B., Huber, R., Tomschy, A., and Jaenicke R., J. Mol. Biol., 246 (1995) 511 Moracci, M., CiarameUa, M., Nucci, R., Pearl, L.H., Sanderson, I., Trincone, A., and Rossi, M., Biocatalysis, 11 (1994) 89 Moracci M., Capalbo L., Ciaramella M. and Rossi M. Protein Eng., 9 (1996) 1191 Pappenberger, G., Schurig, H., and Jaenicke R., J. Mol. Biol., 274 (1997) 676 Perutz, M.F., and Raidt, H., Nature, 255 (1975) 256 Russell, R.J.M., Hough, D.W., Danson, M.J., and Tailor, G.L., Structure, 2, (1994) 1157 Tomschy, A., B6hm, G., and Jaenicke R., Protein Eng., 7 (1994) 1471 Yip, K.S.P., Stillman, T.J., Britton, K.L., Artymiuk, P.J., Baker, P.J., Sedelnikova, S.E., Engel, P.C., Pasquo, A., Chiaraluce, R., Consalvi, V., Scandurra, R., and Rice, D.W., Structure, 3 (1995) 1147 Weismann, C., Beste, G., Hengstenberg, W., and Schultz, G.E., Structure, 3 (1995) 961
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
317
A strategy for engineering thermostability: the case of cyclodextrin glycosyltransferase Joost C.M. Uitdehaag and Bauke W. Dijkstra* BIOSON Research Institute and Laboratory of Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands. Nowadays, protein engineering strategies to make thermostable enzymes are considered to be dependent on the mechanism of thermo-inactivation. For the enzyme cyclodextrin glycosyltransferase (CGTase), this mechanism is believed to be irreversible unfolding. To reduce the enzyme's inactivation rate, a sensible engineering strategy would be to stabilize flexible loops near the active site. These loops can be detected using the atomic B factors in any of the five CGTase X-ray structures. A comparison of thermolabile and thermostable CGTases shows that Nature has indeed chosen a strategy of stabilizing flexible loops near the active site. On this basis, some site-directed mutagenesis experiments are suggested.
1. I N T R O D U C T I O N
Cyclodextrin glycosyltransferases (CGTases) are enzymes that have the unique ability to produce cyclodextrins (o~(1->4) linked circular oligoglucosides) from starch. CGTases are evolutionary related to o~-amylases, and both enzymes are commonly used during the industrial liquefaction of starch. This is done by jet-cooking at high temperatures to keep the starch soluble, which requires use of thermostable enzymes, like the B. licheniformis o~-amylase I or the Thermoanaerobacter CGTase 2. in contrast, thermolabile CGTases have been isolated with very interesting specificities, like the production of a single size of cyclodextrin 2. To take advantage of such a specificity in an industrial context, thermostability will have to be engineered into these thermolabile CGTases through site-directed mutagenesis. * corresponding author
318
This is a challenging task, which requires foremost detailed insight into the enzyme's individual structure and function 3. Fortunately, for CGTases, five different three-dimensional structures have been solved 48, with sequence homologies > 60%, of which two are thermostable CGTases 6,7. These structures were extensively compared by Knegtel et al. 7 in combination with biochemical data, which identified many sites contributing to thermostability. Below we will provide reasons for focusing on two sites in particular, and suggest mutations to enhance thermostability in CGTases. From this, the consequences for other (~-amylase like proteins will be discussed.
2. SITES OF THERMOLABILITY
The proper approach to stabilize an enzyme depends on whether the thermoinactivation process is reversible or irreversible 9. In the case of reversible inactivation, there is an equilibrium between functional (folded) and unfunctional (unfolded) enzyme. Consequently, engineering thermostability means shifting this equilibrium towards the functional state by either stabilizing the folded state or destabilizing the unfolded state. This has been the focus of numerous strategies in the past 3'9'1~ However, in the case of (x-amylases, and thus probably also for the related CGTases, thermo-inactivation proceeds by irreversible unfolding 11, which means that once the enzyme has unfolded, it can never become functional again. Here thermostability is determined by the rate of unfolding, which depends on the free energy difference between the folded state and the transition state for unfolding 9. To decrease the unfolding rate this energy d i f f e r e n c e has to be increased. Therefore, a protein engineering approach should aim at making structural differences between folded and transition states as high in energy as possible. In contrast, it is ineffective to modify sites in the protein that have a similar structure in folded and transition state. The structures of folded and transition states are probably very alike. This is indicated by the fact that unfolding is often a highly cooperative process 3, meaning that its transition state comes early in the unfolding route. Therefore, structural differences between folded and transition state are likely to be confined to the sites where the unfolding begins. These are the loops that are structurally not very well stabilized (flexible). Moreover, unfolding of the loops closest to the active site would
319
have a direct effect on the catalytic process. Therefore, the flexible loops near the active site should be our engineering target in CGTase. Identifying flexible loops can be done using the atomic temperature factors (B factors) that are incorporated in any X-ray structure determination of sufficient high resolution. Such a B factor is directly related to the fluctuations around the average position of an atom 12, and correlates strongly with the calculated entropy of a loop13. Analyses of B factors, however, in studies of stability, are sparse14.
3. COMPARING
FLEXIBILITY IN CGTASES
To identify the flexible loops near the active site in CGTases, we first had to define the 'active site'. On the basis of a structure of a CGTase-maltononaose complex 15, we concluded that residues 70-270 and 310-350 (from a total of 686, using Bacillus circulans strain 251 CGTase numbering) were in or near the active site. Subsequently, for the various CGTases, equivalent C(~-positions were found from the pairwise superpositions of their X-ray structures. The B factors of those Ce~ atoms were taken to represent the local flexibility. If a particular amino acid position was deleted in a structure, it was assigned a B factor of 0.01 A 2, to account for deletion as a tactic to reduce flexibility. Subsequently, all B factors were divided by the average B factor of their structure, to enable comparisons. The CGTases were divided into two groups, thermostable (thermophile)" B. stearothermophilus 6 (Bstea) and Thermoanaerobacterium 7 (Tabium ) and thermolabile (mesophile): B. circulans strain 2515 (Bc251), B. circulans strain 84 (Bc8) and B. sp. 10118 (consisting of two independent molecules). Within each group, the normalized B factors were averaged. The thermolabile B factors were plotted versus the thermostable B factors in figure 1. For most residues, there is no consistent difference in B factor between thermostable and thermolabile CGTases: these dots lie close to the diagonal in figure 1. Some dots lie above the diagonal, indicating residues that are more flexible in thermostable CGTases, but these are all non-active site residues (white circles). In contrast, all positions near the active site that have consistently high B factors in thermolabile CGTases (grey circles on the right in figure 1) lie below the diagonal, and are thus stabilized in the thermostable CGTases. This shows that Nature has indeed targeted the flexible loops around the active site, which comprise residues 89-92 and 335-336. These residues were also mentioned by Knegtel et aL7 as part of the loops 88-95 and 335-339, which can now be singled
320
out as probable prime determinants of thermostability. Interestingly, another study of CGTases 16 located thermostability in the first and second N-terminal domains, where also the two loops 88-95 and 335-339 are located.
Figure 1. The average normalized B factors (see text) of the C(~ atoms in three thermolabile (x-axis) and two thermostable (y-axis) CGTase structures. Each circle represents one equivalent amino acid position in all five superimposed CGTase structures. A grey circle indicates a residue near the active site, a white circle a residue distant from the active site, residues mentioned in the text are numbered. A position with a high x-coordinate or a high y-coordinate signifies a consistent large flexibility in thermolabile or thermostable CGTases respectively. The diagonal is drawn for reference; residues deviating from the diagonal were targeted by evolution towards thermostability. The labelled residues were checked manually for involvement in crystal contacts in all CGTase structures. Omitting these crystal contacts from the averaging procedure and redrawing the figure still showed the special positions of the labelled residues.
321
4. MAKING
MUTATIONS
The loops 88-95 and 335-339 were discussed by Knegtei et aL 7 Loop 88-95 forms substrate binding site +315, and it was concluded that it was stabilized in Tabium CGTase compared to Bc251 CGTase by hydrophobic interactions attained through insertion of an extra residue at position 92. In addition an Asn 88 Pro substitution might reduce entropy of the unfolded loop in the transition state, in accordance with the 'proline rule '1~ In Bstea CGTase, the 88-95 loop is one residue shorter than in Bc251 CGTase, and residue Asp 89 augments the internal hydrogen bonding network, which is very loose in Bc251 CGTase. To stabilize the 88-95 loop by protein engineering, one can envisage to transfer the entire sequence from a thermostable to thermolabile CGTase, including surrounding residues that interact with it. Alternatively, point mutations could be rationally devised to stabilize the loop. For example in Bc251 CGTase, a mutation Y89D could improve internal hydrogen bonding, deletion of residues 89-91 is possible, or a S90P mutation could reduce flexibility.
Figure 2: Structure of the loop comprising residues 330-344 in diverse CGTases: Bc251 (white), Bstea (grey) and Tabium (black). Indicated are individual residues contributing to loop stability. The picture was made with MOLSCRIPT 17.
322
The other flexible loop near the active site, with residues 335-339, is close to the important catalytic residue Asp 328, and therefore its instability could directly interfere with catalysis. Knegtel et aL7 found that the loop 335-339 is one residue shorter in the thermophile Tabium CGTase and that it is more stabilized by hydrogen bonds and hydrophobic contacts, compared to the mesophile Bc251 CGTase (see figure 2). Interestingly, again a proline is present (Pro 341), which reduces the flexibility in the Tabium loop, whereas none of the thermolabile CGTases have Pro in the loop 335-339. In the thermophile Bstea CGTase, residues 335-339 are also stabilized by a proline, Pro 339, in addition to a Lys 376Asp 335 salt bridge (figure 2). Again, a mutational strategy to stabilize this loop can involve mimicking nature, by for instance introducing in Bc251 CGTase the R339P and K341P mutations, or by deleting residue Asn 335. However, due to sequence differences in the 335-339 loop, point mutations might have unexpected effects, and modelling and crystallographic techniques will have to be used to relate the biochemical behaviour to the structure. Fortunately, the speed of modern day protein crystallography is such, that it can easily be used as an analytical tool to study mutations.
5.
CONCLUSIONS
The wealth of structural information available about CGTases has enabled us to understand the structural basis of thermostability in this enzyme class 7. If thermoinactivation proceeds via irreversible unfolding, as is the case for CGTases, the flexible loops around the active site should be the first targets for a stability engineering strategy. This notion is supported by comparing B factors in five mesophile and thermophile CGTase structures, where Nature appears to have targeted loops 88-95 and 335-339. Stabilization of these loops has occurred through a promiscuous use of salt-bridges, proline substitutions, residue deletions and hydrophobic interactions, all factors which have been implicated in thermostability 9,1~ This shows that not the type of interaction is essential, but its strength and position, which depends on an enzyme's individual three-dimensional structure. Therefore, the results for CGTases cannot straightforwardly be applied to related enzymes with limited sequence homology, like the e~-amylases. However, if denaturation by irreversible unfolding is the general mechanism for all enzymes from the (z-amylase family 11'18, flexible loops near the active site might be a
323
common target to enhance thermostability. It should be noted that two known stabilizing mutations in B. licheniformis (~-amylase, H133Y and A209V 18 are not positioned in regions of high flexibility. The structure of these mutations is however not known, and their influence on flexibility is therefore uncertain. Currently we are working on CGTase mutants to test our theories, and rationalize the phenomenon of thermostability in this enzyme class. ACKNOWLEDGEMENTS
The authors thank Dr. Finn Drablos for providing a CGTase and e~-amylase sequence alignment. Conny Uitdehaag and Prof. Lubbert Dijkhuizen are thanked for their comments on the manuscript. REFERENCES
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.
Machius, M., Wiegand, G. & Huber, R. J. MoL BioL 246, 545-559 (1995). Pedersen, S., Dijkhuizen, L., Dijkstra, B.W., Jensen, B.F. & Jorgensen, S.T. Chemtech 19-25 (1995). Jaenicke, R. Eur. J. Biochem. :202, 715-728 (1991). Klein, C. & Schulz, G.E.J. MoL BioL 217, 737-750 (1991). Lawson, C.L., van Montfort, R., Strokopytov, B., Rozeboom, H.J., Kalk, K.H., de Vries, G., Penninga, D., Dijkhuizen, L. & Dijkstra, B.W.J. Mol. BioL 236, 590600 (1994). Kubota, M., Matsuura, Y., Sakai, S. & Katsube, Y. Denpun Kagaku 38, 141-146 (1991). Knegtel, R.M.A., Wind, R.D., Rozeboom, H.J., Kalk, K.H., Buitelaar, R.M., Dijkhuizen, L. & Dijkstra, B.W.J. MoL BioL :256, 611-622 (1996). Harata, K., Haga, K., Nakamura, A., Aoyagi, M. & Yamane, K. Acta Cryst. i:)52, 1136-1145 (1996). Shaw, A. & Bott, R. Curr. Opin. Struct. BioL 6, 546-550 (1996). Watanabe, K., Masuda, T., Ohashi, H., Mihara, H. & Suzuki, Y. Eur. J. Biochem. 226, 277-283 (1994). Tomazic, S.J. & Klibanov, A.M.J. BioL Chem. 263, 3086-3091 (1988). Drenth, J. Principles of protein X-ray crystallography (Springer-Verlag, New York, 1994). Meirovitch, H. & Hendrickson, T.F. Proteins Struct. Funct. Genet. 29, 127-140 (1997). Fontana, A. How nature engineers protein (thermo)stability. in Life Under Extreme Conditions (eds. di Prisco, G.) 89-113 (Springer-Verlag, Berlin, 1991). Strokopytov, B., Knegtel, R.M.A., Penninga, D., Rozeboom, H.J., Kalk, K.H., Dijkhuizen, L. & Dijkstra, B.W. Biochemistry 35, 4241-4249 (1996). Kaneko, T., Song, K.-B., Hamamoto, T., Kudo, T. & Horikoshi, K. J. Gen. Microbiol. 135, 3447-3457 (1989). Kraulis, P.J.J. AppL Crystallogr. 24, 946-950 (1991). Declerck, N., Machius, M., Chambert, R., Wiegand, G., Huber, R. & Gaillardin, C. Prot. Eng. 10, 541-549 (1997).
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
325
Thermophilic esterases and the amino acid "traffic rule" in the h o r m o n e sensitive lipase subfamily Giuseppe Manco*, Ferdinando Febbraio* and Mos~ Rossi *~ *Institute of Protein Biochemistry and Enzymology, CNR, via Marconi 10, 80125, Naples, Italy *University of Naples "Federico II", via Mezzocannone 16, 80100, Naples, Italy
We recently reported on the identification, over-expression and partial characterisation of a new esterase from the thermoacidophilic eubacterium Bacillus acidocaldarius homologous to the hormone sensitive lipase (HSL)-like sub-group of the esterase/lipase family. A comparative analysis of the amino acid compositions in this family quite unexpectedly revealed some trends in amino acid changes that are presumably related to both protein thermal stability and/or thermophilicity. Here we further extended this analysis to an acetylesterase (AES) of E.coli and to a new homologous gene from the hyperthermophilic archaea Archeoglobus fulgidus. Moreover an analysis of data with the statistical approach of Argos et al.(1979) confirmed the aforementioned trends in amino acid changes going from psychrophilic to hyperthermophilic members of the HSL group and revealed some new features.
1. INTRODUCTION Some studies (1, 2) have suggested that esterases, lipases and cholinesterases belong to a large family of phylogenetically related proteins with representatives in the domains of Eukarya and Bacteria and include proteins lacking enzymatic activity. Three subfamilies have been identified: the C group includes cholinesterases from Vertebrate and Invertebrate, lipases from Fungi, a number of esterases and some non-enzymatic proteins; the L group includes lipases from Vertebrate and Bacteria, lipoprotein lipases, lecithin-cholesterol acyltransferases and related non-enzymatic vitellogenins from flies; the third group, H, was also named HSL by Hemil~i et al. (1) since these authors reported the cloning and sequencing of a gene from B. acidocaldarius which encoded a protein of unknown function (ORF3) homologous to the HSL from human and rat. In this group were included also: the N-acetylphosphinothricintripeptide deacetylase from Streptomyces hygroscopicus; the acetylhydrolase from Streptomyces viridochromogenes; the lipase 2 from Moraxella TA144 (MOL); an esterase from Acinetobacter calcoaceticus (ACE); the esterase AES encoded by the ORF203 of E. coli; Vsh5, a protein from the slime mold Dictiostelium discoideum; and the recently sequenced human liver arylacetamide deacetylase (HDAC). Unfortunately, it is unclear at the moment whether this sequence similarity implies comparable biochemical properties for these enzymes. New enzymes from thermophilic bacteria could add new insights into the evolutive
326 relationships in the esterase/lipase family from one hand and from the other they could allow fundamental studies in the field of protein stability for this class of proteins. Moreover they are giving rise to growing interest in the biotransformations field. To date, carboxylesterases have been purified and partially charaeterised only from Sulfolobus acidocaldarius (3), Bacillus stearothermophilus (4-6) and B. acidocaldarius (7). We recently reported on the purification of a new esterase from B. acidocaldarius and demonstrated its identity with the aforementioned ORF3 (8). The gene was over-expressed in E. coli purified and partially characterised to get more insight into the structure-function relationship of this interesting representative of the HSL group (9). Here we report the alignment of an hyphotetical protein from A. fulgidus (Accession number: AEOOO782 from GenBank TM database, Release 103), with several members of the HSL group and an analysis of the respective amino acid compositions with the aim to identify amino acid changes that are related to both protein thermal stability and/or thermophilicity.
2. MATERIAL AND METHODS
2.1. Computer analysis The procedure for the multisequenees alignment was as previously described (8). In short the alignment between Candida rugosa lipase (CRL), Torpedo californica acetylcolinesterase (TAC) and Geotrichum candidum lipase (GCL) based on 3-D superpositions was taken from ref. (10). EST2, AFES, and MOL sequences, automatically aligned by the MAXHOM (11) method of the PHI) server (http:www.embl-heidelberg.de) were manually aligned to the above three sequences on the basis of secondary structure correspondences. Secondary structure predictions were obtained using the PredictProtein PHDsec program (12) from the PHD server. For HSL the secondary structure-driven alignment to CRL, TAC and GCL was taken from ref. (13) although the protein sequence was from mouse, which displays a 27.3 % identity with EST2 in a 289 amino acid overlap using the FastA program of the GCGSequence analysis package (14). Gaps were kept to minimum and, where possible, were not allowed inside the structural elements. 2.2. Amino acid composition analyses The amino acid composition analyses were all obtained from the respective published sequences by using the AAeomp program (SBDS). 2.3. Statistical analysis The statistical analysis of aligned sequences was perfomed according to Argos et al. (15) by generating an "ad hoe" program writed in Q-basic language on a Hewlett Packard personal computer. 3. RESULTS AND DISCUSSION In a previous report (8) we predicted for EST2, a carboxylesterase from the thermoacidophilic eubacterium B. acidocaldarius encoded from the ORF3 gene (1), the or/13 hydrolase fold of the esterase/lipase family. Moreover we made a secondary structure-driven multisequence alignment of EST2 with other members of the HSL subfamily.
b3
CRL
(2 8)
- FAEPP
.... VGNLRFKD.
TAC
(3 5)
- FAEPP
.... VGNMFRR
MOL
(8 5 )
-LAIDPK.
GCL EST2
HSLM AFES
CRL
(2 9) (I)
(253)
(4)
- FADPPVG.
.D L R F K H P Q
. .P V P Y S G S L D G Q K F T L Y G P L C M Q Q N P E G T Y E E N L P K A A L D L V M O S
VSRLLSL..
MVWI YGGAFVFGSSASYPGNGYVKESVEM
EST2
LVY.____~GGGWVVGDLETH.
MOL
.QHERVERVEDRT
. .D V Y N G K Y L A Y T E
MLFFHGGGFCIGDIDTH.
. .D P V C R V L A K D G .
.G Q P V V F V S
.E V V L V S L S Y R V G A F G F L . .W A V V S V D Y R M A P
VVHIHGGGFVAQTSKSH.
CRL
TYKGKPLFRAGIMOSGAMVPSDA.
. . VDGIYGNEI
. . .S R D L F R R A I L Q S G S P N C P W A .
.S V S V A E G R R R R A V E L G R N L N C N L N
C S I E s H . . .D A L C R R I A R L S
LVYYHGGGFVI
.V P I I S I D Y S L A P
86 GCL TAC
EST2
MOL HSLM AFES
CRL GCL TAC
EST2
MOR
HSLM AFES
TYNGKKLFHSAI
LQSGGPLPYFDSTSVGPESAYA
. . .V R V P D G I M A A .
YPVTDYEAEYP YPVTTLQ
.... EDFI KHQILIYPVVNF.
QLCPSYIVVAEL.
DILRDEG.
NLPPALIITAEY.
DPLRDEG.
Figure 1:
.............
EGYLLTGGMMLWFRDOYLNSL
.............
EGLWI
FPELSGRKP
LDQKIMSWFSEQYFSR
[38
" u I,., FLS KQLSGLPVLGTFHSNDIVFODYLLGSG
DPMLDDS.
KLYAEALNKAGVKVEIENFE
VMFARRLRDLGQPVTL~E
EVFGQMLRRAGVEASIVRYR
.L R I D P S
. NTPGF.
.S D E E L I H C L R E K K P Q E L I D V E W N V L P F D S I F R F S
.S L L E F G
LAYAELLQKEGVQVQTYTVL
" us., FGESAGSMSVMCHIL
...............
...............
...............
...............
.............
...... DLIHGFAQFYS__~L. SPG .......... HQGLGNQTTYI
...... GVLHGFINYYPV.
LKA ..........
...... DLPHGFLSLAALCRET.RQATEFCVORI
WNDGDN
.I FGESAGAMSVAHQLVAYGGDN
..
FGESAGGASVGMHILSPG.
KI FVGGDSAGGNLAAAVS
IMARDSG
. . LAYSSLRLSYLPRPDGVNITDDMYALVEGKY
. 9. F G L L P Q F L G F G P R P D G N I .......
QKTT.
I PDAAYELYSGRY
FVPVIDGEFFPTSLESMLNGNF
EELTHPWFSPVLYP
HSGLPQSHPLI
SVMHG
..............
u2 ,,, SLIYNNAFIAFA.
DLS
..............
DNT
FA ..............
DLE
S PTAESVRPTESMRRspLLAPDSM
EEDKFNPLASVI
............. SA. YRRYFI SFAN. YT. AEEEALSRRIMHYWATFAKTG-
...... GAPHGFINLMSV.
V
13r.4 ,~
GCDASASDNETLACLRSKSSDVLHSAQNSYDLKDL
...... ASSWLNS
..........
~3,~
. KLACLRGVS.SDTLFDATN.
EGLLLDHNDAEVFNSAYTQ
S 1 GVMGLVQRDTSLFLRDLRLG
VAPTP
...... RSRAL
.L G A S P S R I V L S G D S A G G C L A A L v t F K N S L A D
EHKFPAAVYDCYDATKWVAENAEE
.............
YPA
DEAA
D .F H L D P A R I A V G G D S A G G N L A A V T . S I L A K E R G
EYPAPTALKDCLAAYAWLAEHSQS
DEGT I LAPVAINATTTPnvNRWTYLATQLHNLPFLGTFHGSDLLFQYYAGPWS DEGSFFL... LYGAPGFSkgTYLYFFNHRASneWMGVIHGYEIEFVFGLPFVKELN.
GLPPAYIATAQYYD.PLRDVG. SLPPVHIVACAL_.
..........
.F G G D P K T V T I
KPDSP
KSTTV
........
EAPFPRALEECFFAYCWAVKHCDDLLGSTGERICLAGDSAGGNLCITVSLRAAAYG
0:6, ~ NA.. G C . G S A S D .
.FGGDPKVM
EHKFPAAVEDAYDALQWIAERAA
. .S W E L Y G
[37 " u I~., u 2~,, A N I IP _V _I~ N N D E G T F F G T s r E Y F K O L F n g T K Y S AKVPYITGNQE. KKTQILLGVNK.
FDLL~.
VRVYQQ.
135 FGGDPTKVTI
SQEAPGNVGLLDQRMALQWVHDNIQF
............
l~1 7
. . .G P A Y I A F _ _ Q _ ~ Y P S T G Y D P A H P P A S I E E N A
. . .L P R P L A Q L P L .
ALHG.
............
.N S T V V S V D Y R L A P
IKGRNGDIR
...................
TAEGNTNAGLHDQRKGLEWVSDNIAN
............
HSLM
AFES
.G G D A I
............
RAVVFSVDYRLAP
.H E F C H ~ C A Q T G .
.E p Y L K S W A O E L G .
INYRTGPYGFL
........
SWQDKTIANADGGDMTVRCYQSTQNSERKST.
%., .I KAEGSANAGLKDQRLGMQWVADNIAA.
84 GKPI.IHVSVNYRVSSWGFLAGDE
MDLPGRTLKVRMYRPEGVEPP
...... LARLISYDLREGQDSKVLNSLAKSEGPRLDVRPRPHQAP
9P P E A F E M P L T S D P R L T V T I S P P L A H T G P A P V
a,,, 83 MLWIFGGGFEVGGTSTFPPAQMITKSIAM.
EMWNPNREMSEDCLYLNIWVPSPRP
...............................
-MPIDPVYYQLAEYFDSLpfSSAREYREAINRIYEERNRQLS.
...... ANLPV
...... DAKLPV
GPLyqGSVSMNEDCLYLNVFRPAGTKP
....................
9. P A P D Y K H L S A Q Q F R S Q Q S L F P P V K K E P V A E V R E F D
.L R Q K F G T D A V S L ~ P S V W Q Q N A D A S G S T E N A V
MVWIYGGGFYSGSSTL.
...............
.... PEPKKPWSGVWNASTYPNNCQQYVDEQFPGFSGS
GCL
TAC
82 ..... KVFEAVLPLSEDCLTINVVRPPGTKAG
..... PFTGSYQGLKANDFSSACMQLDPGNAISLLDKVVGLGKILPDNLR.
MPLDPVIQOVLDOLNPd!4__.
-SLANMAS...
c%,,2
9. L K
TD-
H-
ATKALVRIAEKLRDALA
INEFACLVQNLLTSEGDKPNLRA RLI LTPPAAPLN
ARDAINQIAALLVFD
Multiple sequences alignment of AFES with some HSL subfamily enzymes and with CRL, TAC and GCL. The SwissProt codes are: CRL, p20261; GCL, p04058; TAC, G406508; MOL, p24484; HSLM, p54310; The sequence of AFES, was taken from GenBank TM database, Release 103 Accession number: AEOOO782, whereas that of EST2 was taken from ref. (1). [3-strands and or-helices of CRL (16) are indicated as single or double lines, respectively. The stretches of hypothetical [3-strands or a-helices in EST2 (8) and HSL (13), estimated using the PredictProtein PHDsec program, are indicated in the same way. The serine, aspartic and histidine residues, most likely to be at the active site in the HSL-group, are indicated by an asterisk. The residue couples in lowercase letters indicated insertions that are not shown. Finally, the residues which are conserved among all sequeces or in the HSL group are indicated in bold.
328 Searching the GenBank TM database we have identified a new hypothetical protein from the hyperthermophilic sulphate-reducing archaeon A. fulgidus reported to share 42% identity with the A. calcoaceticus esterase, a member of the HSL group. In Figure 1 we show a multisequence alignment which includes the new sequence. The alignment revealed the presence of highly conserved residues already identified by Cygler et al. (10) in the esterase/lipase family, as well as residues which are common to the HSL group. It is worth mentioning that the region corresponding to the lid in CRL (16) was partially deleted in the AFES sequence. This finding, suggests that the enzyme is not a lipase. The location of the elements belonging to the catalytic triad Ser...Glu/Asp...His proved particularly interesting. Table 1 Amino acid compositions comparison ASE AMINO MOL ACE (37~ ~ ACIDS (5~ ~' (30~ e'
Ala Cys Asp Glu Phe Gly His Ile Lys Leu Met Asn Pro Gin Arg Ser Thr Val Trp Tyr
# mole% 50 11.55 8 1.85 26 6.00 19 4.39 9 2.08 26 6.00 17 3.93 19 4.39 21 4.85 52 12.01 6 1.39 21 4.85 25 5.77 25 5.77 13 3.00 26 6.00 25 5.77 25 5.77 6 1.39 14 3.23
HSL* (37~ a'
mole% # mole% # mole% # 10.32 28 8.70 32 10.03 80 9 2.82 14 1.81 (2.19) 4 1.24 6.58 31 4.00 21 6.52 21 5.33 48 6.19 (5.47) 16 4.97 17 4.70 33 4.26 15 4.66 15 6.58 58 7.48 16 4.97 21 6 1.88 18 2.32 10 3.11 3.76 22 2.84 19 5.90 12 8 2.51 16 2.06 16 4.97 13.81 45 13.98 37 11.60 107 3.13 19 2.45 7 2.17 10 8 2.51 19 2.45 (2.19) 10 3.11 6.27 55 7.10 (6.72) 23 7.14 20 5.64 24 3.10 19 5.90 18 5.64 56 7.23 13 4.04 18 4.08 72 9.29 17 5.28 13 5.96 41 5.29 (5.31) 17 5.28 19 4.08 40 5.16 13 4.04 13 5 1.57 6 0.77 3 0.93 5.33 16 2.06 10 3.11 17
EST2 (65~ a'
AFES (83oc)e'
# mole% # mole% 37 11.94 28 9.00 1 0.32 3 0.96 21 6.77 22 7.07 20 6.45 24 7.72 12 3.87 16 5.14 22 7.10 17 5.47 8 2.58 6 1.93 11 3.55 23 7.40 12 3.87 10 3.22 34 10.97 25 8.04 6 1.94 5 1.61 6 1.94 11 3.54 28 9.03 16 5.14 12 3.87 11 3.54 16 5.16 24 7.72 13 4.19 20 6.43 8 2.58 5 1.61 23 7.42 25 8.04 4 1.29 3 0.96 16 5.16 17 5.47
Amino acid compositions were all obtained from the respective published sequences. * In the case of HSL the results do not change significantly (data in parenthesis) if the phosphorylation domain [1] is excluded from computation. a, This is the temperature midpoint of source microorganism. The location of the active serine (Ser 160) is clearly indicated by the conserved pattern "GlyXISX2Gly" and its topological position in a 13-eSer-a motif aider strand 135 (Fig. 1) (17). In the C-terminal part of the protein, His 285, which was well conserved in all members of the HSL subfamily (HGF/A motif) as well as in CRL, TAC and GCL, is most likely to be the
329 histidine of the active site. The acidic element of the triad could be Asp 255 according to the alignment with the EST2 sequence (8). Site-directed mutagenesis experiments recently made on the correspondig Asp and His residues in human HSL (13) and in EST2 (G. Manco, unpublished results) are in agreement with this assignment Thus the HSL subfamily includes hydrolases from Bacteria, Archaea and Eukarya, as well as from psychrophilic, mesophilic, thermophilic and hyperthermophilic sources. Comparative studies within this group would be of valuable interest in the field of protein stability determinants. We previously observed (9) some trends of amino acid changes in the HSL subfamily going from the psychrophilic to the thermophilic enzymes. Now we have extended this analysis to the AES and AFES enzymes. In Table 1 we show the amino acid compositions of several members of the HSL group. As a result some trends in amino acid changes were still evident (Glu and Cys) some other were not confirmed (Pro and Asn) and finally a tendency to change Thr was more evident. Note that the Cys content in the AFES was higher than in EST2. A possible explanation for this behaviour is that being A. fulgidus an anaerobic archaeon the enzyme is much less sensitive to Cys oxidation. With the aim to analyse more carefully these results we decided to exploit the method of Argos et al.(15) which should be able to test statistically the significance of the amino acids drifts towards the "warmer" sequences. A program able to analyse automatically a multisequence alignment was generated and used to analyse the sequences indicated in Table 1. The multisequence alignment (not shown) was based on the same criteria as those used for the alignment of Figure.1. The results obtained are shown in Table 2 and diagrammatically in Figure 2. The outpout of the program is a matrix of the ratios Eij between the net weighted exchange per degree Celsius to the standard error. Only values of Eij_>_l.7 c were taken into account and ranked from the higher to the lower. Table 2 Representative amino acid changes Exchanges with Cold ~ Hot Ei] _>1.70 L Q L S T T Q Q E
=~ =~ =~ =~ =~ =~ ~ ~ =~
V P A A L R K R A
5'82 2.97 2.28 2.23 2.19 2.16 2.16 2.05 2.04
Cold
L E L C Q G N S
~
Hot
~ ~ ~ ~ =~ =~ ~ =~
Y R E L N R I P
Exchanges with Eij >_1.70
1.89 1.88 1.88 1.80 1.77 1.72 1.72 1.70
First of all it is clear that the observed changes are not coincident with those reported from Argos et a/.(15) except for the Ser--~Ala change in the cold--hot direction. However we confirm the existence of "psichrophilic" and "thermophilic" residues with Gly, Ser, and Gin belonging to the former and Ala, Arg, Leu, Glu and Asn belonging to the latter. In contrast with the Argos data on the "cold" side we also found Leu, Thr, Glu, Asn and Cys whereas on the "hot" side we have Val, Pro, Ile, Leu, Lys, and Tyr. The difference in results could be attributed to the higher temperature range spanned by the sequences we used and/or to the
330
1 4.-.
N ~._.. Q . . . ~
P 4--
S -.~
Y
Figure 2. The observed exchanges in the direction of thermophilic preferences. The marked arrows indicate that a single mutation event in the codon was necessary for the amino acid change. higher difference in the evolutive distances between the tested species. Alternatively differences could be family-specific. Much more enzyme families need to be analysed to answer this point. As shown in Figure 2 several of found exchanges could be easily obtained by means of single base change per codon. Other exchanges, especially those involving Leu require additive mutational events. A possible explanation could be the higher Leu content of all sequences respect AFES. REFERENCES
1. Hemil~i,H., Koivula, T.T., and Palva, I. (1994) Biochim. Biophys. Acta 1210, 249-253. 2. Krejei, E., Duval, N., Chatonnet, A., Vincens, P., and Massouli~, J. (1991) Proc. Natl. Acad. Sci. USA 88, 6647-6651. 3. Sobek, H. and Gorish, H. (1989) Biochem. J. 250, 453-458. 4. Matsunaga, A., Koyama, N., and Nosoh, Y. (1974) Arch. Biochem. Biophys. 160, 504513. 5. Owusu, R. K., and Cowan, D. A. (1991) Enzyme Microb. Technol. 13, 158-163 6. Wood, A. N., Femandez-Lafuente, R., and Cowan, D. A. (1995) Enzyme Microb. Technol. 17, 816-825. 7. Manco, G., Di Gennaro, S., De Rosa, M., and Rossi M. (1994) Eur. J. Biochem. 221,965972. 8. Maneo, G. Adinolfi, E., Pisani, F.M., Carratore, V., and Rossi, M. (1997) Protein and Peptide Letters 4, 375-382. 9. Manco, G. Adinolfi, E., Pisani, F.M.,Ottolina,G., Carrea, G. and Rossi, M. (1998) Biochem. d. in press. 10. Cygler, M., Schrag, J. D., Sussman, J. L., Harel, M., Silman, I., Gentry, M. K., and Doctor, B. P. (1993) Protein Science 2, 366-382. 11. Sander C. and Schneider, R. (1991) Proteins Struct. Funct. Genet. 9: 56-58. 12. Rost, B., and Sander, C. (1993) I Mol. Biol. 232, 584-539. 13. Contreras, J.A., Karlsson M., Osterlund, T., Laurell, H., Svensson, A., and Holm, C. (1996) J. Biol. Chem., 271, 31426-31430. 14. Devereux, J., Haeberli, P., and Smithies, O. (1984) Nucleic Acids Res. 12, 387-395. 15. Argos, P, Rossmann, M.G., Grau, U.M., Zuber, H., Frank, G., and Tratschin, J.D. (1979) Biochemistry, 18, 5698-5703. 16. Grochulski, P., Li, Y., Sehrag, J.D., BouthiUier, F., Smith, P., Harrison, D., Rubin, B., and Cygler, M. (1993)I Biol. Chem. 268,12843-12847. 17. Derewenda, Z.S. and Derewenda, U. (1991) Biochem. Cell. Biol. 69, 842-851.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
331
Cloning and stabilization of NAD-dependent formate dehydrogenase from C a n d i d a boidinii by site-directed mutagenesis H. Slusarczyk, M. Pohl and M.-R. Kula Institut ~ r Enzymtechnologie, Heinrich-Heine Universitat Duesseldorf, Forschungszentrum Juelich, D-52426 Juelich, Germany
The gene of the NAD-dependent formate dehydrogenase from the yeast Candida boidinii was cloned by polymerase chain reaction and expressed in E. coil The recombinant enzyme was stabilized by site-directed mutagenesis. Two cysteine residues probably located at the surface of the protein were exchanged against various aliphatic amino acids and the mutant enzymes were tested for stability against oxidation by air in the presence of catalytic amounts of Cu(II). All mutants were significantly more stable against oxidative stress than the wildtype enzyme. Additionally, the mutant enzymes were characterized with respect to their temperature optima, kinetic constants and activation energy of the enzymatic reaction. The results of these studies indicated that the catalytic properties of the mutant enzymes have not been altered by the mutagenesis compared to the wildtype enzyme.
1. INTRODUCTION NAD-dependent formate dehydrogenase (EC 1.2.1.2) is found in many methylotrophic bacteria and yeasts [ 1]. The enzyme catalyzes the oxidation of formate to carbon dioxide with the concomitant reduction of NAD to NADH. Because of the favourable thermodynamic equilibrium of the reaction the enzyme is used for in situ regeneration of NADH during asymmetric synthesis of chiral compounds [2,3].The enzyme is efficiently formed as an intracellular enzyme by the yeast Candida boidinii, if methanol is the only energy and carbon source available [4].In order to perform enzymatic reductions on large scale a production and downstream process has been established [5]. Though FDH from C. boidinii (Cb-FDH) can be prepared in large quantities and was used in several enzymatic redox reactions [2,3] the enzyme is less stable against oxidative stress [6] resulting in higher costs for the biocatalyst in synthesis reactions. Therefore, the enzyme from the yeast was cloned and expressed in E. coli. This paper deals with the stabilization of the recombinant enzyme by site-directed mutagenesis and the enzymatic characterization of the resulting mutants. Their properties are compared to the native enzyme from C. boidinii.
332 2. MATERIALS AND METHODS
2.1 Cloning of the fdh-gene The cloning procedure is described elsewhere [7] 2.2 Site-directed mutagenesis In vitro site-directed mutagenesis was performed according to the method of rio et al. [8] using pUC-FDH as a template. 2.3 Expression in E coli Recombinant bacteria were cultivated on LB-medium containing 100 ~tg AmpiciUin/ml. Expression was induced by addition of 1 mM IPTG (isopropyl-13-D-thiogalactoside) aider A550 reached 0.5. The cells were harvested by centrifugation 3 h ~ e r induction. Cell desintegration was carried out by glass beads in a mixer mill and the cell free crude extract were analyzed for activity and protein content according to the method of Bradford [9], using BSA as a standard. 2.4 Measurement of FDH acitivity FDH activity was assayed spectrophotometricaUy at 340 nm using a Shimadzu photometer UV160. The assay mixture was composed of 162 mM sodium formate and 1.62 mM NAD in 100 mM potassium phosphate buffer, pH 7.5. The mixture was thermostated to 30~ unless otherwise stated and the reaction was started by addition of limiting amounts of enzyme. The increase in absorbance at 340 nm was followed and activities calculated by using e = 62201 mol1 cmq for NADH at 340 rim. One unit is defined as the amount of enzyme which catalyzes the reduction of 1 ~mol NAD per min at pH 7.5 and 30~ 2.5 Kinetic constants Kinetic constants were derived from duplicate measurements of initial velocity under conditions varying one substrate and saturating concentrations of the second substrate. The corresponding data were fitted to a model for double substrate kinetics according to MichaelisMenten, using the software 'Scientist for Windows'(Microsofl| 2.6 Temperature dependencies of enzyme activity Temperature dependencies of the activity of the native, the recombinant wildtype enzyme (rec-FDH) and the mutant enzymes have been tested by increasing the incubation temperature under standard assay conditions. Activation energies were calculated from Arrhenius plots using the linear segments of the lnvmax versus 1/T plots. 2.7 Thermal inactivation Thermal inactivation experiments were carried out by incubating the native enzyme in 100 mM potassium phosphate, pH 7.5, over a temperature range between 30 an 70~ After 20 rain aliquots were removed and the residual activity and protein content were determined. 2.8 Stability in presence of Cu(IO All enzymes were tested for stability against oxidative stress by incubation in the presence of 10 ~M CuCI2 in 100 mM potassium phosphate, pH 7.5, at 25~ Controls were incubated without CuC12. At certain time intervalls, aliquots were removed and residual activity was determined.
333 3. RESULTS AND DISCUSSION
The fdh gene from Candida boidinii was amplified from a genomic DNA template by polymerase chain reaction and cloned into pUC 18. The Cb-FDH open reading frame (ORF) codes for a 364-aa protein (40 370 Da) which shows highest homology to FDH from C. methylica [10]. Only a single aa exchange could be detected: Leu at position 87 in Cb-FDH corresponds to Ser in Cm-FDH. The wt-fdh gene was cloned into the expression vector pBTac2 and transformed into E. coli JM105. FDH was overproduced after induction with IPTG to about 20-25 % ofE. coli soluble cell protein. The thermal stability of Cb-FDH was tested by stepwise increasing the incubation temperature up to 70~ At several time intervals aliquots were removed and subsequently the residual activity and the soluble protein content were determined. Fifty percent of activity was lost after incubation at 57~ for 20 min. Similar values have been obtained for FDHs from the yeasts Kloeckera sp.(55~ and Pichia pastoris (54~ [ 11]. Thermal inactivation of the CbFDH above 55~ is paralleled by a loss of soluble protein in the enzyme preparation. The loss of activity at higher temperatures is caused by an aggregation process, which is not readily reversible lowering the temperature again. Kelly and Zydney [12] examined the role of oxidation of sulfhydryl groups in bovine serum albumin (BSA) on fouling during microfiltration. They found that aggregation of BSA during the filtration process could be minimized by adding sulfhydryl groups-modifying agents like iodacetamide or dithiothreitol to a prefiltered protein solution. The results presented in their study demonstrated that B SA aggregation in these filtration systems occurred via an intermolecular thiol-disulfide interchange reaction, possibly in combination with thiol oxidation reactions. Based on this observations two cysteine residues, Cys23 and Cys262 in the recombinant wtFDH (rec-FDH) were exchanged by various aliphatic amino acids by site-directed mutagenesis resulting in four different mutants. Using the cfistal structure of FDH from the bacterium Pseudomonas sp. 10] [ 13] as a model, the substituted cysteine residues are likely to be located at the surface of the protein and therefore highly accessible for oxidation by molecular oxygen. The stability of the mutant enzymes in comparison with the wt-enzyme against oxidative stress was estimated by incubation of partially purified enzyme preparations in the presence of catalytic amounts of divalent copper ions. Traces of divalent ions like Cu(II) or Fe(II) were shown to promote oxidation processes at thiol groups[ 14,15].
334
Figure 1. Residual activities ofwt-FDHs and various mutant enzymes incubated in 100 mM potassium phosphate, pH 7.5, for 15 h at 25~ in absence and presence of 10 ~tM CuCI2, respectively.
All mutant enzymes showed a significantly higher stability in presence of CuCI2 compared to Cb-FDH as well as rec-FDH (Figure 1). Even after 15 hours of incubation in presence of Cu(II) ions a residual activity of about 70% could be determined in all mutant enzyme samples whereas the wt-activity decreased to about 8%. The results suggest that the inactivation of the wt-enzymes (Cb-FDH and rec-FDtO is caused by oxidation of free sulfhydryl residues. Further studies characterizing the inactivation process are under investigation. In order to compare the catalytic and enzymatic properties of the mutant enzymes and the wt-enzymes, all enzymes were characterized with respect to their temperature optima, kinetic constants and activation energy, respectively. Figure 2 demonstrates that both the wt-enzymes and the mutant enzymes show broad temperature optima between 45 - 55~ Obviously, the temperature optima of the mutant enzymes are slightly reduced to 50~ compared to the wtenzymes. This effect is probably due to a decrease in conformational stability of the mutant enzymes caused by the amino acid substitutions.
335
100 --A--C23S -- x7-- C262V - o - C23SC262V - +- C23SC262A /
80 o~" 60 "~
,\ i ~
~~
40 20
10
'
20'
'
3'0
'
' ;o 40' temperature [~
'
60'
'
70'
Figure 2. Effects of temperature on the activity of Cb-FDH, rec-FDH and various mutant enzymes. The linear segments of the Arrhenius plots are used for determinating the activation energies. In all enzyme preparations FDH-activity is proprotional to the incubation temperature in the range of 20 - 40~ and the calculated activation energies are similar or increased compared to the wt-enzyme (Table 1). Table 1 Kinetic constants and activation energies of Cb-FDH, rec-FDH and various mutant enzymes. Km Formate [mM]
Km HAD [pM]
Ki NADH [BM]
activation energy [kJ/mol]
Cb-FDH
5.6 +/- 0.4
45 +/- 2.9
17 +/- 1.2
55
recFDH
5.9 +/- 0.4
37 +/- 3.0
21 +/- 1.8
54
FDH-C23S
5.5 +/- 0.4
44 +/- 3.5
16 +/- 1.3
62
FDH-C262V
4.3 +/-0.5
41 +/-5.2
8.8 +/-1.2
55
FDH-C23S/C262V
4.5 +/- 0.6
44 +/- 5.6
11 +/- 1.5
59
FDH-C23S/C262A
5.0 +/-0.3
45 +/-3.0
13 +/- 0.9
68
As demonstrated in Table 1, the kinetic constants of the mutant enzymes compared to the wtenzymes are not significantly altered.
336 The present study represents the succesful stabilization of NAD-dependent formate dehydrogenase from C. boidinii by site-directed mutagenesis of two oxidation-sensible cysteine residues. These amino acid exchanges did not influence the catalytic properties of the enzyme. The potential of the mutant enzymes as stable biocatalysts for the in situ regeneration of NADH is under investigation.
ACKNOWLEDGEMENT This work was supported by BMBF AZ 03D0031 C7.
REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.
V.O. Popov and V.S. Lamzin, Biochem. J. 130 (1994) 625. A.S. Bommarius, K. Drauz, W. Hummel, M.-R. Kula and C. Wandrey, Biocatalysis 10 (1994) 37. G. Krix, A.S. Bommarius, K. Drauz, M. Kottenhahn, M. Schwarm, M.-R. Kula, J. Biotechnol. 53 (1997) 29. H. Sahm, Adv. Biochem. Eng. 6 (1977) 77. D. Weuster-Botz, H. Paschold, B. Striegel, H. Gieren, M.-R. Kula, and C. Wandrey, Chem. Eng. Technol. 17 (1994) 131. H. Sch0tte, J. Flossdorf, H. Sahm and M.-R. Kula, Eur. J. Biochem. 62 (1976) 151. H. Slusarczyk, PhD Thesis, D0sseldorf University, Germany, 1997. S.N.Ho, H.D. Hunt, R.M. Horton, J.K. Pullen and L.R. Pease, Gene 77 (1989) 51. M.M. Bradford, Anal. Biochem. 72 (1976) 248. S.J. Allen and J. Holbrook, Gene 162 (1995) 99. J.J. AUais, A. Louktibi and J. Baratti, Agile. Biol. Chem. 47 (1983) 2547. S.T. Kelly and A.L. Zydney, Biotechnol. Bioeng. 44 (1994) 972. V.S. Lamzin, Z. Dauter, V.O. Popov, E.H. Harutyunyan and K.S. Wilson, J. Mol. Biol. 236 (1994) 759. J.M. Kolthoff and B.R. Willeford jr., J. Am. Chem. Soc. 80 (1958) 5673. L. Michaelis, J. Biol. Chem. 84 (1929) 777.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
U s e f u l n e s s of b a c t e r i a l t h e r m o s t a b l e analysis
337
e n z y m e s in clinical chemical
K. Tomita a, K. Nomura ~, T. Miura" and H. Shibata b aCollege of Engineering, Kanto Gakuin University, 4834 Mutsuura, Kanazawa-ku, Yokohama 236-8501, Japan bHead Office, Iatron Laboratories, Inc., 1-11-4 Higashikanda, Chiyoda-ku, Tokyo 101-0031, Japan
Since enzymes have many excellent characteristics, e.g., high substrate and reaction specificities, they have become popular in clinical chemical analysis. However, many of the usual enzymes tend to lose their activities on storage, especially in a solution state, even at rather low temperature, which limits their uses remarkably. On the other hand, enzymes from thermophiles are known to be superior to usual enzymes in thermal and storage stabilities. The author et al. [1] have successfully prepared many thermostable enzymes from a thermophilic bacterium, Bacillus stearothermophilus (BS). Moreover, we have obtained several enzymes from a mesophilic bacterium, Zymomonas mobilis (ZM), and found they also have good thermal stability[2]. In this paper, we show their usefulness as tools in clinical chemical analysis, with focusing the application of two kinds of them, i.e., glucokinase (GlcK, EC 2.7.1.2) and alanine dehydrogenase (AlaDH, EC 1.4.1.1), to the determination of some components in a serum. 1. MATERIAL AND METHODS 1.1. Material Glck was prepared from BS cell or ZM cell. AlaDH was prepared from Escherichia coll C 600 cell carrying the plasmid-encoding BS AlaDH gene [3]. Glucose 6-phosphate dehydrogenase (G6PDH, EC 1.1.1.49, from Leuconostoc mesenteroides) was obtained from Boehringer Mannheim GmbH, Mannheim, Germany. The other chemicals were commercial products of analytical grade. 1.2. Methoda The enzymatic activities of GlcK and AlaDH were measured at 30~ on the bases of the change in absorbance at 340 nm due to the formation of NADPH (GlcK coupled with G6PDH) or NADH (AlaDH) according to the usual method.
338
Four kinds of reagents for the determination of the components in biological fluids, mainly sera, were compounded from adequate reactants mentioned in the Sections of 2.3.-2.6. A sample was added to the reagent, incubated at 30~ (glucose determination) or 37~ (the others), and the formation of NADPH or NADH was spectrophotometricaUy measured at 340 nm. 2. RESULTS AND DISCUSSION 2.1. Properties of GlcK The main properties of BS GlcK [4] and ZM GlcK [2] were as follows. The molecular weights were determined to be 68,000 (BS) and 65,000 (ZM), and 32,000 (BS) and 33,000 (ZM) for the subunits. The optimal pHs for the enzymatic activities were 8.5 (BS) and 7.0-8.0 (ZM). BS GlcK was stable in a wide pH range over 8, and ZM GlcK was stable in a pH range from 6.0 to 8.0. Noteworthily, both GlcKs were fully active even after treatment for 60 rain at 60~ (BS) and 50-55~ in the buffer solution, whereas hexokinase (HK, EC 2.7.1.1) from yeast rapidly lost its activity at temperature above 35~ (Figure 1).
100 75 50 9~
25
~
0 0
25
50
75
100
Temperature (~
Figure 1. Thermal stabilities of both GlcKs dissolved in the buffer solution. Comparison with yeast HK. After incubation for 15 rain at different temperatures, the remaining enzymatic actim'ties were measured. 0 : BS GlcK; A: ZM GlcK; m: yeast HK.
Both GlcKs showed no activities when fructose, mannose and glucosamine were used as phosphoryl acceptors. On the other hand, both the mammarian GlcK and the yeast HK were fully active to these monosaccharides. The Km values were
339 determined to be 0.1 mM (BS) and 0.2 mM (ZM) for glucose and 0.05 mM (BS) and 0.5 mM (ZM) for ATP, being much lower than those of mammarian GlcK and the yeast HK. 2.2. Properties of AlaDH The molecular weight was determined to be 230,000, and 38,000 for the subunit. The optimal pH for the enzymatic activity was 10.4. AlaDH was stable over a wide pH range above 7, moreover, over a wide temperature range. AlaDH was fully active even after treatment for 60 rain at 70~ in the buffer solution. AlaDH showed no activity when L-leucine, L-isoleucine and L-valine were used as hydrogen donors. The Km values were determined to be 10.0 mM for L-alanine and 0.26 mM for NAD § These properties were similar to other AlaDH from mesophilic Bacillus sp., except for pH and thermal stabilities.
2.3. Determination of glucose In clinical chemical analysis, the determination of glucose is a very important factor for the diagnosis of diabetes mellitus. Although the method using HK as the key enzyme (HK method) is recommended as the reference method for glucose determination by several organizations, e.g., American Association for Clinical Chemistry[5], the properties of HK are much inferior to those of GlcK as described above. The application of BS GlcK to the glucose determination (GlcK method) [6] was
1t,511tl
2500 N 211110 o 15110
5OO 0
10
20
30
40
Time(day) Figure 2. Storage stability of the reagent solution for the determination of glucose. Comparison of GlcK method and HK method. After storage for several ten days at room temperature, each reagent solution was used for the determination of glucose in a serum sample. O: GlcK method; m: HK method.
340 attempted according to the principle 9Glucose is phosphorylated by GlcK in the presence of ATP to yield glucose 6-phosphate (G6P); G6P is converted to 6phosphogluconate (6PG) by G6PDH in the presence of NADP, which results in the formation of NADPH; finally, glucose is determined as the change in absorbance at 340 nm. The assay had a high degree of precision, i.e., coefficient of variation (CV) was 0.7-1.3% (within-run reproducibility) and 1.1-2.3% (between-run reproducibility ), and was linear up to about 8000 mg 1"1. A good correlation was observed between GleK method and HK method for both plasma and urine samples. The outstanding superiority of GlcK method to HK method was the storage stability of the reagent solution. The solution based on GlcK method was found to be stable at least for about one month at room temperature (Figure 2) [7] and 21 months at 10~ [8] , whereas that based on HK method was stable for only a few days at room temperature (Figure 2) [7]. ZM GlcK was also applied to glucose determination [2], and gave as excellent results as BS GlcK. As for the stability of the reagent solution, ZM GlcK gave a little inferior results to BS GlcK, but much superior to HK. 2.4. Determination of creatine kin~ae activity The determination of creatine kinase (CK, EC 2.7.3.2 ) activity is much in use for the diagnosis of myocardial infarction or progressive muscular dystrophy. The current method based on using HK (HK method) involves the same disadvantage in the stability of the reagent solution as that shown above. The new method [9] was also developed by using BS GlcK based on the principle: CK acts on creatine phosphate in the presence of ADP to yield ATP; followed by two reactions coupled with GlcK and G6PD, respectively, ATP causes a formation of NADH. The assay had a high degree of precision, i.e., CV was 1.12.2% (within-run reproducibility), and was linear up to about 2000 U 1-'. It was noteworthy that the reagent solution was found to be stable at least for one week at room temperature and for 13 months at 10 ~ [8]. The conventional HK method gave only one day stability at room temperature. 2.5. Determination of magnesium ion The determination of magnesium ion is a matter of significance for the diagnosis of renal diseases and gastrointestinal disorders. The colorimetric method, in which Xylidyl Blue is utilized as a detector of magnesium ion, is currently used, but this method has a defect in specificity. The atomic absorption spectrophotometry is looked upon as the most reliable method, but requires an expensive instrument. The reagent for magnesium ion determination [10] was successftflly composed with using GlcK on the basis of the paper of Tabata et a]. [11]. The principle of the assay was based on the fact that magnesium ion-ATP complex acts as a substrate of GlcK, as shown in Figure 3. The concentration of magnesium ion is then determined as an increasing rate in absorbance at 340 nm due to NADPH turnover. This simple and rapid spectrophotometric method was found to have a good correlation with the atomic
341 absorption spectrophotometry and to be very little affected by several cations in a serum such as K § Na § Zn s+, Ca s§ NH4+ and so on. The present method was recognized as to be more specific to magnesium ion than the Xylidyl Blue method. The assay had a high degree of precision, i.e., CV was 0.85-1.7% (within-run reproducibility) and 1.25-2.15% (between-run reproducibility), and was linear up to about 100 mg 1-'. This linearity range was considerably wider than that of the Xylidyl Blue method. The present reagent had an excellent stability at least for about one month at 10~ in liquid form. It should be emphasized that this method is the world's first practical enzymatic method for measuring metal ion. GlcK Glucose
~
Mg- ATP ATP
~
G6P M g - ADP Mga+ ~
""~ADP
G6PDH G6P + NADP +
6PG + NADPH + H §
Figure 3. The schematic principle of GlcK method for the determination of magnesium ion.
2.6. Determination of,/-glu~myltransferase activity Determination of~/-glutamyltransferase (v .GT, EC 2.3.2.2) activity is routinely employed in the diagnosis of hepatic diseases. The colorimetrlc method, in which ~/-glutamyl 4-nitroanilide (,/-Glu4NA) is utilized as a substrate for ,/-glutamyl donor, is most commonly used in current, but ~/-Glu4NA has a serious problem, i.e., poor solubility in water. The present method [12] was ultraviolet spectrophotometric one, in which the readily soluble substrate ~/-glutamyl dipeptide was used as the ~/-glutamyl donor and thermostable amino acid dehydrogenase as the attxiliary enzyme. In the preliminary experiment, ~/-glutamylalanine (~/-GluAla) was found to be the most specific and sensitive donor substrate. This method is consisted of two enzymatic reactions: First, ~/-GluAla releases L-alanine by the action of~/-GT in the presence of glycylglycine(GlyGly); second, L-alanine is converted to pyruvate by AlaDH in the presence of NAD, accompanied by NADH formation. NADH is continuously monitored at 340 nm. The present method is strictly stoichiometric, since one NADH is produced from one ~/-GluAla by every ~/-GT action. The assay had a high degree of precision, i.e., CV was 0.8-2.8% (within-run reproducibility) and about 2.2% (between-run reproducibility), and was linear up to about 700 U 1-1. The present method well correlated to the ,/-Glu4NA method. Although this method had about one-third
342 of the sensitivity to the T -Glu4NA method, it was adequate for the practical assay. The storage of the reagent solution did not affect the measuring activity at least over about 30 days at 10~ 3. CONCLUSIONS The bacterial thermostable enzymes, GlcK and AlaDH, had high degrees of stability, especially thermal and storage stab'dities, which made them useful in clinical chemical analysis. The other properties were also well suited for the analysis, i.e., low Km values, high substrate specificities and so on. These enzymes were successfully applied to the establishment of new enzymatic methods which gave long term stability of the reagent solutions as well as rapid, accurate and reliable results.
REFERENCES 1. K.Tomita, K.Nagata and K.Okada, Ann. N.Y. Acad. Sci., 672 (1992) 178. 2. K.Tomita and K.Nomura, Ann. N.Y. Acad. Sci., 750 (1995) 338. 3. Y.Sakamoto, S.Nagata, N.Esaki. H.Tanaka and K.Soda, J. Ferment. Bioeng., 69 (1990) 154. 4. S.Kamei, K.Tomita, K.Nagata, H.Okuno, T.Shiraishi, A.Motoyama, A. Ohkubo and M.Yamanaka, J. Clin. Biochem. Nutr., 3 (1987)1. 5. P.Duncan, J.Neese, D.Bayse and C.Stewart, Clin. Chem., 20 (1974) 882. 6. K.Tomita, S.Kamei, K.Nagata, H.Okuno, T.Shiraishi, A.Motoyama, A. Ohkubo and M.Yamanaka, J. Clin. Biochem. Nutr,. 3 (1987) 11. 7. K.Tomita, K.Nagata, H.Kondo, T.Shiraishi, H, Tsubota, H.Suzuki and H.Ochi, Ann. N. Y. Acad. Sci., 613 (1990) 421. 8. K.Tomita, K.Nomura, H.Kondo, K.Nagata and H.Tsubota, J. Pharm.Biomed. Anal., 13 (1995) 477. 9. H.Kondo, T.Shiraishi, M.Kageyama, K.Nagata and K.Tomita, J.Clin.Biochem. Nutr., 3 (1987) 17 10. T. Shiraishi, H. Suzuki, H.Ochi, K. Kawahara, H. Kondo, K. Nagata and K.Tomita, Jap. J. Clin. Chem., (in Japanese) 20 (1991) 37. 11. M.Tabata, T.Kido, M.Totani and T.Murachi, Agric. Biol. Chem., 50 (1986) 1909. 12. H.Kondo, M.Hashimoto, K.Nagata, K.Tomita and H.Tsubota, Clin. Chim. Acta, 207 (1992) 1.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
343
Effect of temperature and Ca 2+ on the degree of multiple attack exhibited by mesophilic and thermophilic a - a m y l a s e s B. Kramhofi and B. Svensson Carlsberg Laboratory, Department of Chemistry, Gamle Carlsberg Vej 10, DK-2500 Copenhagen Valby, Denmark. The activity of (z-amylases depends on Ca 2§ and temperature. In the starch hydrolysis these depolymerases exert a distinct number of catalytic events per enzyme-substrate encounter referred to as the degree of multiple attack. The present results indicate that the degree of multiple attack for the thermophilic a-amylase from Bacillus licheniformis is reduced from about 2 to 1 following an increase in the Ca2+-concentration from 0.1 mM to 5 mM. The degree of multiple attack is unchanged following an increase in the temperature from 37 ~ to 80 ~ at both Ca2§ The multiple attack by an (z-amylase from a related thermophilic alkalophilic Bacillus is reduced at the lower temperature and in low Ca 2§ but is unaffected by the temperature at the higher Ca2§ At 37 ~ and at the low Ca 2+concentration a reduced multiple attack was exhibited by the mesophilic a-amylase from porcine pancreas and isozyme 2 from germinated barley seeds. In contrast, barley a-amylase 1 maintained the level of multiple attack found at the high Ca2§ 1. INTRODUCTION The degree of multiple attack exhibited by a-amylases is defined as the average number of catalytic events following cleavage of the first glycosidic bond during the life-time of an individual enzyme-polysaccharide complex [1]. The degree of multiple attack for a given enzyme thus contributes to the understanding of the nature of the productive enzyme-substrate interaction. Using a substrate with a high degree of polymerization, the degree of multiple attack for a number of a-amylases was previously determined to vary from 1.9 for the oramylase from Aspergillus oryzae to 6 for porcine pancreatic a-amylase [ 1]. The aim of the present study was to determine the role of Ca 2§ and of temperature in the degree of multiple attack exhibited by a number of thermophilic and mesophilic a-amylases having extended substrate binding sites. The chosen a-amylases from bacteria and higher plants all have an active site containing 9-10 consecutive binding subsites each accommodating one substrate glucosyl residue [2-5]. In contrast, pancreatic a-amylase, which is known to exert a high degree of multiple attack, has an active site comprising only 5 subsites [6]. Ca2. is required for activity of all a-amylases [7-9]. The different three-dimensional structures have a common Ca2+-site that stabilizes a small domain containing side chains essential for the substrate binding by bridging this domain and the catalytic (13/or)s-barrel domain. This characteristic domain interplay occurs at the subsites in the direction of the nonreducing end from the bond to be cleaved [4,10,11 ]. Various additional Ca2"-sites exist in B.
344
fichemformis and barley a-amylases [4,10].
In the multiple attack mechanism the nonreducing end product of the hydrolysis is envisaged to reorganize in the substrate binding cleft for subsequent catalytic events.
2. METHODS The enzymatic digests were performed with amylose (Sigma) as substrate at a concentration of 0.1% in 2 % DMSO in the appropriate buffer. The degree of polymerization (DP) of the amylose was 440 determined as described by Robyt and French [1]. In the case of the thermophillic Bacillus and the barley cx-amylases, the buffer was 20 mM Na-acetate, pH 5.5 with either 0.1 mM or 5 mM CaCI2. For porcine pancreatic a-amylase 20 mM MES, pH 6.9 was used with CaC12 added as described above. Enzymatic digests were performed in a small scale. Samples (1 ml) from a 10 ml reaction mixture of substrate with enzyme were removed at intervals during the initial phase of substrate degradation. Hydrolysis of substrate was monitored after iodine staining as the decrease in "blue value" measured at 620 nm [ 1,12]. In separate experiments it was found that the initial stage of substrate hydrolysis corresponds to a decrease in blue value of about 20%. This was usually obtained within 20-45 min after addition of the enzymes (data not shown). The Bacillus enzymes were kind girls from Novo Nordisk, Denmark. The barley a-amylase isozymes were prepared in-house as described earlier [5]. Porcine pancreatic o~-amylase was from Sigma. "l~he amounts of enzyme used in the digests were as follows: Bacillus licheniformis a-amylase: 10-30 mU/ml; a-amylase from alkalophilic Bacillus: 30-100 mU/ml; barley a-amylase,isozyme 1 (AMY1): 25-70 mU/ml and isozyme 2 (AMY2): 10-25 mU/ml. One unit is defined as the amount of enzyme that causes an increase in absorbance of 1.0 at 620 nm within 15min at 37 ~ using insoluble Blue Starch (Pharmacia) as a substrate [9,13]. Porcine pancreatic a-amylase was used at a concentration of 5-10 mU/ml, where 1 unit, according to Sigmae is defined as the amount of enzyme liberating 1 mg of maltose from starch at pH 6.9 and 20 ~ The enzyme action was arrested by addition of trichloroacetic acid to a final concentration of 0.1 mM. The. remaining polymeric amylose was subsequently precipitated with 67% ethanol [ 1], and lefi: overnight at 4 ~ The contents of reducing sugar in the total digest (RVt) and of the ethanol precipitate (RVv) were determined by the copper-bicinchoninate method adapted to a microtiter plate assay using glucose as a standard [14]. The degree of multiple attack is defined as (RVt/RVv)-I as described previously [1], and was calculated using analytical data obtained from several samples from each digest with blue values in the range of 80-90% of the initial value. 3. RESULTS AND DISCUSSION
3.1. Thermophilic a-amylases The degrees of multiple attack were determined for two bacterial thermophilic a-amylases at two different temperatures and at two different Ca2+-concentrations (Table 1). Significant degrees of multiple attack, in the range 1-3, are seen for both Bacillus a-amylases. This means that during each encounter of enzyme with the substrate, in average 1-3 glycosidic bonds are hydrolyzed in addition to the first cleaved bond. The a-amylase from B. licheniformis has a slightly reduced degree of multiple attack in 5 mM CaCI2 compared to the value obtained at
345 the low CaCl2-concentration. The specific activity of the a-amylase from B. licheniformis, measured with insoluble Blue Starch as substrate, increases somewhat but not dramatically as a function of Ca 2§ in the present concentration range [3]. The combined observations are difficult to interpret in relation to the three-dimensional structure of the enzyme, where three Ca2§ and one Na+-ion by crystallography are found to stabilize the active conformation [4]. Finally, it is shown that an increase in the temperature of the enzymatic digests from 37 ~ to 80 ~ does not affect the degree of multiple attack, suggesting that the stability of the enzyme-substrate complex is not temperature-dependent (Table 1). Table 1 De~rees of multiple attack exhibited by thermophilic a-amylases 0.1 mM CaCI2
5 mM CaCI2
Enzyme source
37 ~
80 ~
37 ~
80 ~
B. licheniformis
2.2
2.1
1.2
1.5
Alkalophilic Bacillus
1.9
2.6
2.6
2.5
The degree of multiple attack exhibited by the alkalophilic Bacillus ct-amylase tends to be reduced at the low Ca 2+ -concentration, but only at 37 ~ This suggests that at this Ca 2§ concentration the stability of the enzyme-substrate complex is temperature-dependent as well as Ca2+-dependent. At 5 mM CaCI2 the temperature of the enzymatic digest has no effect on the degree of multiple attack, perhaps reflecting a more efficient stabilization of the structural integrity at the higher concentration of CaC12.
3.2. Mesophilic or-amylases The degrees of multiple attack were determined for mesophilic enzymes from a higher plant (barley) and from an animal source (porcine pancreas), respectively (Table 2). All values are obtained from digests performed at 37 ~ Table 2 Degrees of multiple attack in mesophilic a-amylases Enzyme
0.1 mM CaCI2
5 mM CaCI2
Barley AMY1
1.4
1.1
Barley AMY2
0.5
1.2
Porcine pancreatic or-amylase
4.7
5.9
The degree of multiple attack exerted by both barley a-amylase isozymes is lower than for the bacterial and mammalian enzymes. In the case of the AMY1 isozyme, reduction of the
346 Ca2+-concentration of the digest does not significantly change the degree of multiple attack. In contrast, the degree of multiple attack of the AMY2 isozyme is strongly CaZ§ and has the lowest value (0.5) in the present study in 0.1 mM CaCI2 (Table 2). This may be a consequence of the lower affinity for Ca2+ of AMY2 compared to AMYI [8]. The effect of Ca2§ on the degree of multiple attack observed with the barley or-amylase isozymes parallels the Ca2§ of the activity of these enzymes towards insoluble Blue Starch. The activity of AMY1 is almost Ca2+-independent in the used concentration range, whereas the specific activity of AMY2 is strongly reduced at low Ca2§ and has a maximum at 5-15 mM CaClz [ 13]. Both isozymes contain structural Ca2+[ 10], but how the present results can be related to the three-dimensional structures of the enzymes remains to be elucidated. Interestingly, the ligands for the three Ca2§ bound in barley a-amylase are conserved between AMY1 and AMY2 [ 10]. Since the degree of multiple attack represents an average estimate and is defined as the number of bonds hydrolyzed after the first bond cleavage in each enzyme-substrate encounter, the result obtained with AMY2 (Table 2) indicates that at a low Ca2§ AMY2 acts on amylose by multiple attack as well as by the multichain mechanism. The multichain mechanism is the classical random action, where the enzyme hydrolyzes only one bond per productive encounter with the substrate [ 1]. As previously reported porcine pancreatic a-amylase exhibits a high degree of multiple attack.[ 1]. This is confirmed in the present study, where a value close to 6 was found (Table 2). The degree of multiple attack determined for pancreatic a-amylase using the ethanol precipitation technique adapted to small scale digests agrees well with previous results obtained using substrates with DP values ranging from 17 to 1000 [ 1,15,16]. Thus, the degree of multiple attack found for porcine pancreatic a-amylase is several fold greater than that exhibited by the thermophilic bacterial as well as the mesophilic plant a-amylases in the present study. It is tempting to speculate that this property might be related to the fact that porcine pancreatic a-amylase possesses a shorter active site with only 5 subsites [ 11] compared to the 9-10 subsites long binding sites found in the other a-amylases used in this study [2-6]. Also in the case of the porcine pancreatic or-amylase some dependence on the Ca2+-concentration is observed (Table 2). This agrees with the finding that the enzyme molecule contains the structural Ca2§ found in all the other a-amylases [4,10,11 ]. The increasing insight into the role of Ca2+ and other ions on the functional conformation of the different a-amylases from B. licheniformis, barley and porcine pancreas [4,10,11,17] renders it highly relevant to assess the influence of Ca2+ on enzymatic properties including the mechanism of multiple attack for which the structural basis is presently poorly understood. ACKNOWLEDGMENT The kind gift of the Bacillus enzymes by Henrik Bisg~rd-Frantzen (Novo Nordisk, Denmark) is gratefully acknowledged. REFERENCES
1. J.F. Robyt and D. French, Arch. Biochem. Biophys. 122 (1967) 8. 2. H. Outtrup, H. Bisg~,rd-Frantzen, P. Rahbek Ostergaard, M. Dolberg Rasmussen and P. van der Zee, Patent no. PCT WO 95/26397. 3. D. Tull, H. BisgS,rd-Frantzen and B. Svensson, in preparation.
347 4. M. Machius, N. Declerck, R. Huber and G. Wiegand, Structure 6 (1998) 281. 5. E.H. Ajandouz, J. Abe, B. Svensson and G. Marchis-Mouren, Biochim. Biophys. Acta 1159 (1992) 1932. 6. E.H. Ajandouz and G. Marchis-Mouren, Carbohydr. Res. 268 (1995) 267. 7. B.L. Vallee, E.A. Stein, W.N. Summerwell and E.H. Fisher, J. Biol. Chem. 234 (1959) 2901. 8. D.S. Bush, L. Sticher, R. van Huystee, D. Wagner and R.L. Jones, J. Biol. Chem. 264 (1989) 19392. 9. I. Matsui and B. Svensson, J. Biol. Chem. 272 (1997) 22456. 10. A. Kadziola, J.-i. Abe, B. Svensson and R. Haser, J. Mol. Biol. 239 (1994) 104. 11. M. Qian, R. Haser, G. Buisson, E. Du~e and F. Payan, Biochemistry 33 (1994) 6284. 12. J.M. Bailey and W.J. Whelan, J. Biol. Chem. 236 (1961) 969. 13. K.W. Rodenburg, N. Juge, X.-J. Guo, M. Sogaard, J.-C. Chaix and B. Svensson, Eur. J. Biochem. 221 (1994) 277. 14. J.D. Fox and J.F.Robyt, Anal. Biochem. 195 (1991) 93. 15. A.K. Mazur and H. Nakatani, Arch. Biochem. Biophys. 306 (1993) 29. 16. H. Kondo, H. Nakatani, K. Hiromi, R. Matsuno, J. Biochem. 84 (1978) 403. 17. A. Kadziola, M. Sogaard, B. Svensson and R. Haser, J. Mol. Biol. (1998) in press.
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 1998 Elsevier Science B.V.
349
Degradation and denaturation of stable enzymes Dion Thompson a, Roberto Fem/mdez-Lafuente b, Cesar Mateo c, Don A. Cowan b, Jose M. Guisan r and Roy Daniel a. Thermophile Research Unit, University of Waikato Private Bag 3105, Hamilton, New Zealand. a
b Department of Biochemistry University College London, United Kingdom. c Departamento de biocat/disis. Instituto de Catalisis. CSIC, Campus Universidad Autonoma 28049 Madrid, Spare.
1. INTRODUCTION We are interested in the mechanisms which govern the upper temperature limit for enzyme activity. The loss of enzymic activity at high temperatures can be separated into two broad classes. The first is the loss of tertiary and secondary structures. The second is the loss of primary structure through covalent modifications or cleavage. These two processes have been referred to as denaturation and degradation respectively I. The aim of the present study is to investigate the relative roles of denaturation and degradation in activity loss in an enzyme system which has reasonable stability at temperatures above 100~ with a view to understanding which of these two processes limits high temperature enzyme stability. The enzyme chosen for study was a xylanase from a stain of Thermotoga maritima 2. This enzyme has a half life of 5 to 10 minutes at 100~ in the absence of stabilisers. Stabilisation was provided by sorbitol and multi-point covalent immobilisation giving rise to a half life of 2 minutes at 140~ when immobilised and suspended in molten sorbitol.
2. IRREVERSIBLE ACTIVITY LOSS The loss of recoverable activity of an enzyme due to heat denaturation typically follows first order kinetics, eqn(1), with respect to concentration and time, although close examination may revel that second order kinetics provides a marginally better approximation 3.
350 A graph of the logarithm of residual activity against time has a slope of-k. Occasionally, a semi-log graph revels a biphasic behaviour whereupon a sharp transition from a high to a low rate constant is made at some stage through time 2. However, no such behaviour was observed in this work.
d[A] dt where:
-
(~)
-k[A]
[A] = concentration of active enzyme k = the rate constant. t = time
The first order rate constant of xylanase activity loss was determined for a range of temperatures and with different stabilising agents. Figure 1 presents these rate constants as a function of temperature in the form of an Arrhenius plot. The slope of an Arrhenius plot represents the activation energy of the reaction, which in this case is the transition from active to inactive enzyme. High activation energies are characteristic of unfolding of macromolecules while lower values are generally found for covalent modifications (eg ref 4).
In(k) Halflife 4ilsec
tee x34amse noso,t~ I ~
~ 50%sort~ A-----~.-__..,
freexylanase
2
esec
O-
9 41sec \ ~..~.
irm'~lised li ..... n o ~ "'" .... x ~ "-.~.
e,o-2 N 5 mn 9
-. .....
9
,,
tee~l~
-4 !38 rrin I I
-6 5hm -8 341"~ 24
1135
127 I
119 i
111 I
104 I
I
245
25
255
26
265
27
901
oC
275 10001~
Temm~e
Figure 1: Arrhenius plot of the first order rate constants for irreversible loss of activity.
351 Three aspects of the denaturation profiles stand out from Figure 1. First, loss of activity of the free xylanase in low concentrations of sorbitol, <-- 50%, and immobilised xylanase (without sorbitol), show similar activation energies. Secondly, these same samples each show a decrease in activation energy at higher temperatures. Third, the loss of activity of the xylanase in a high sorbitol/low water environment has a significantly lower activation energy. It may be that the steeper slopes represent denaturation through unfolding whereas the flatter slopes show covalent processes predominating. The most stable form of the xylanase, immobilised and suspended in molten sorbitol, has a half life of about 2 minutes at 140~ This is not very different from the results obtained previously using different immobilising procedures on the same enzyme 2 or from those found for the ot-glucosidase from Thermococcus zilligii strain AN1 5
3. METHODS
3.1. E n z y m e s The xylanase was originally purified from Thermotoga maritima strain FjSS3-B. 1 2 and cloned into Escherichia. coli 6 Xylanase assays were performed using 10mM o-nitrophenol-13-xylopyranoside in a pH20 6 phosphate/citrate buffer. 3.2. Heating
Thermostability trials were conducted in flame sealed capillary tubes. 70BL of sample at pHz0 6 was sealed into the tubes before immersing in an oil bath at the required temperature. Cooling was achieved by submerging the tubes into an ice bath. Heating/cooling time for the capillary tubes estimated to be < 4 seconds as found by thermocouple measurement and by the explosion of the occasional tube within 2 seconds of immersion at 190~ (no tubes exploded at 150~ 3.4. Immobilisation 10mg of xylanase was immobilised onto 1g of epoxy-acrylic resin which had undergone activation in the presence of an organic base following the general procedures of ref 7. Immobilised xylanase displayed many of the features typical of immobilised enzymes such as flattened pH/activity profile and general resistance to denaturants (data not shown).
REFERENCES
1. R. M. Daniel, M. Dines and H. H. Petach, Biochemical Joumal, 317 (1996) 1-11 2. H.D Simpson, U.D Haufler and R.M Daniel, Biochemical Journal 277 (1991) 413 - 417 3. B.W. Matthews, H. Nicholson and W.J. Becktel, Proceedings of the National Academy of Science USA, 84 (1987) 6663-6667. 4. S.J. Tomazic and A.M. Klibanov, Journal of Biological Chemistry 263 (1988) No &, 3086-3091 5. K. Piller, R.M. Daniel and H.H. Petach, Biochemica et Biophysica Acta, 1292 (1996) 197-205
352 6. D.J Saul, L.C Williams, R.A Reeves, M.D Gibbs and P.L Bergquist, Applied and Environmental Microbiology, 61 (1995) No 11, 4110 - 4113. 7. R.M. Blanco, J.J. Calvete and J.M. Guisan, Enzyme and Microbial Technology, 11 (1989) 353359
Non-conventional media
a This Page Intentionally Left Blank
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
355
Engineering stability of enzymes in systems with organic solvents Vadim V. Mozhaev ~ b
"Department of Chemistry, Moscow State University,
119899 Moscow, Russia
b Department of Chemical and Biochemical Engineering, University of Iowa, Iowa City, IA 52242, USA 1.
INTRODUCTION
Biocatalysis in organic solvents has become a commonplace. The nonaqueous milieu permits the advantages of organic solvents as the reaction media, such as increased solubility of apolar substrates, shift of thermodynamic equilibrium to favor synthesis over hydrolysis, suppression of many water-dependent side reactions and others, while enzymes provide the opportunity for high selectivity and catalytic rates under mild reaction conditions [1]. Unfortunately, exploiting such advantages is otten limited by the low stability and/or activity of the biocatalysts. Enzymes are known to be denatured in the presence of relatively small amounts of polar solvents [2], and in non-aqueous media the catalytic activity is significantly suppressed in comparison with their aqueous level [3]. The problem of low catalytic activity and stability of enzymes in systems with organic solvents are in the focus of this paper. 2.
ENZYME DENATURATION
Dependence of enzyme activity on concentration of water-miscible solvent in binary mixture with water has a specific character, Fig. 1A [4]. The activity does not significantly
10o .
800 ~ NAo%/~&..a,~ 3
A
8O
a
6O 4O
0
20
40
6O
80
[~voL%
100
0
20
40
60
80
100
Fig. 1. Catalytic activity of r as a function of concentration of organic solvents. A - native enzyme in acetonitrile, B - in ethanol: 1 - native enzyme, 2 - complex of native enzyme with polybrene, 3 - complex with polybrene of the enzyme modified with pyromellitic anhydride. A is the enzyme activity in water-solvent mixture in relation to aqueous activity, Ao.
356 changed in the presence of small solvent amounts but within a narrow concentration range drops down to zero in threshold-like manner then remaining at this zero level [5]. When water content is further decreased to as low as 10-30 vol. %, the activity is regained although being much smaller than that in water. For successful enzyme performance in organic media, several issues should be addressed. Why are enzymes inactivated at small and intermediate co-solvent concentrations and how can enzyme be stabilized under these conditions.9 Such systems with 50-80 vol. % of the polar solvents like dimethyl formamide (DMF) or alcohols are very promising for synthetic needs as they afford excellent solubility of many important compounds like peptides. Also, is it possible to raise enzyme activity in organic media containing only a few percent of water or in neat organic solvent? These systems are interesting due to a possibility of dissolving a vast amount of water-insoluble compounds of non-polar character. And finally, is it possible to develop a universal enzyme catalyst able to effectively work at the concentration range from 0 to 100 vol. % of solvent? 2.1.
Denaturation mechanism
Mechanistic consideration of enzyme inactivation in organic solvents is based on the important prerequisite that the protein molecule is surrounded by a shell formed by water molecules bound to the protein surface by hydrogen bonds. This hydration shell or at least a portion of it represents an integral part of protein structure and is essential for enzyme function [6]. Consequently, displacement of bound water molecules by organic solvent results in a dramatic change of the whole protein structure and leads to denaturation. In line with these ideas, inactivation of enzymes in water-co-solvent mixtures consists of three main steps schematically shown in Fig. 2: i. dehydration of the protein molecule, i.e. removal of a certain critical amount of water molecules from the protein hydration shell; ii. binding of the organic co-solvent by the partially dehydrated protein; iii. conformational transition in the protein molecule resulting in the formation of a denatured form D with concomitant loss of catalytic activity. Thermodynamic consideration of this model led to the following expression for the free energy of protein denaturation (AGo) by organic solvents [7]: AGo = Bo + Bin + B2ET(30) + B3nlogP
(~)
where B0, B l, B2, and B3 are numerical coefficients, which are constant for a given protein regardless of the solvent nature, n is the numerical parameter which depends on the size of cosolvent molecules and gives an idea of how many water molecules can be displaced from the protein surface by one molecule of the organic solvent, ET(30) is the Dimroth-Reichardt parameter, which is directly related to the free energy of the co-solvent solvation, P, the partition coefficient of the solvent in water/octanol biphasic system, is a measure of its hydrophobicity. The values of Er(30), P and n for solvents can be either found or calculated from the data contained in comprehensive handbooks on organic and physical chemistry. Equation 1 has a real predictive force in stating that co-solvents with high values of ET(30) and logP are strong denaturants. In fact, the solvents that are both hydrophobic and have a high solvation capacity, like 1,4-dioxane, tetrahydrofuran (THF), and higher alcohols
357 1. Protein dehydration:
o o + O0
N
b:g
o
2. Binding of solvent by partially dehydrated protein:
m +
k
m
n
m
"---
3. Protein denaturation (conformational transition):
Fig. 2. Schematic presentation of the mechanism of protein denaturation by organic solvents. (e.g., isomers of butanol) cause enzyme inactivation at concentrations as low as 10-30 vol. %. On the other hand, hydrophilic solvents like glycerol, ethylene glycol, and formamide have a small denaturing capacity and at concentrations as high as 50-60 vol. % still do not inactivate many enzymes. Due to this fact, concentrated solutions of these solvents can be a convenient medium for enzyme reactions in binary mixtures of co-solvents with water. Scaling of cosolvents according to their denaturing capacity may have a general character as its correctness has been proved correct in experiments with 8 proteins in more than 30 solvents [7].
2.2.
Stabilization approaches
Based on the model in Fig. 2 one can suggest the ideas for corroborating the enzyme inactivation in organic solvents. Because often inactivation starts with protein unfolding, enzymes can be efficiently stabilized by covalent immobilization. Multi-point attachment to a support affords high eonformational rigidity of the enzyme and consequently high activity may be retained in the presence of significant amounts of organic co-solvents. For example, chymotrypsin attached to polyacrylamide gel with a dozen of covalent bonds is inactivated at 60 vol. % methanol that is by 20 vol. % higher than the native enzyme [8]. Probably due to the same reason, i.e. an increase in rigidity of the protein molecule, also the enzymes cross-linked with bifunctional reagents are very stable to denaturation in water-co-solvent mixtures [9].
358 Multi-point attachment to a support can be also achieved by formation of complexes between proteins and polyelectrolytes, which are stable due to multiple non-covalent, mainly electrostatic bonds. The complexes are prepared in aqueous solution by adding a polycation or polyanion, depending on whether the protein is negatively or positively charged. In systems with an organic solvent, the complexes are not destroyed because electrostatic interactions that support existence of the complex become stronger in media with low dielectrics. Due to multiple interactions with the polyelectrolyte, the enzyme is protected against inactivation by organic solvents and retains catalytic activity at much higher co-solvent concentration [10]. Enzyme protection by polyelectrolytes is totally reversible and is eliminated when the complex is destroyed, for example, by added salt. Oligoamines (spermine, spermidine and others) present another group of compounds that form complexes with proteins due to electrostatic binding. It has been observed that such complexes with oligoamines provide additional stabilization for thermophilic enzymes at high temperature. In complexes with o|igoamines enzymes retain high activity at concentrations of organic solvents which completely destroy the activity of native enzymes. Chymotrypsin has been more effectively stabilized in the complex with the longer spermine than with spermidine, probably due to stronger binding of spermine to the enzyme (Kudryashova et al., submitted). Because electrostatic interactions are so important for stabilization by polyelectrolytes, chym0trypsin was modified with pyromellitic anhydride that afforded introduction of several dozen of additional negatively charged carboxylic groups, and the complex with polycation was further obtained. In this complex catalytic activity of the modified enzyme was retained over a much broader range of ethanol concentrations (curve 3 in Fig. IB) than the activity of nonmodified enzyme in free state (curve 1) and in the complex with polybrene (curve 2) [11]. Apparently, complexes of modified enzymes with polyelectrolytes are good candidates to be considered as universal biocatalysts in systems with different concentrations of organic solvent. Because complexes with polyelectrolytes have proved to be so effective in stabilizing enzymes in organic solvents, it was a challenge to use them in such a difficult case as media rich in DMF. This is an ideal solvent for many reactions as it provides a high solubility for many compounds; however, DMF has a bad reputation because it inactivates irreversibly many enzymes. Inactivation of chymotrypsin by DMF was reduced in the complex with polyanion that enabled us, probably for the first time, to carry out peptide synthesis in the presence of as high as 60 vol. % of DMF and as low as 6 vol. % of water, the rest being acetonitrile (Vakurov et al., submitted). Under such a harsh condition, the native enzyme was almost inactive. Enzyme denaturation by organic solvents includes dehydration of protein molecule as an important step of the process (Fig. 2). One can assume that proteins will be less amenable to this denaturation in case their hydration shell is tightly hold due to stronger interaction of water molecules with the functional groups on the protein surface. In fact, introduction of polar and charged groups into the protein molecule by using covalent modification increases affinity of protein to water: much more water is bound by modified chymotrypsin especially in the region of water sorption isotherm corresponding to hydration of protein polar groups (Rees et al., submitted). These modified enzymes retain catalytic activity in the presence of solvent concentrations that induce inactivation of the native enzymes. Such stabilization has been observed for chymotrypsin modified with pyromellitic anhydride, glyceraldehyde [12] and polyethylene glycol [13]. Most probably, the newly attached carboxylic and hydroxylic groups attract water more efficiently, thus retarding protein dehydration caused by organic solvents. It is not excluded that similar mechanism, i.e. strong attraction of essential water molecules, is
359 achieved when amino acid residues on the protein surface are replaced by using site-directed or random mutagenesis [14, 15]. Another stabilization approach recently developed by Arnold [ 16] and Wong [ 17] suggests to increase rigidity of the protein molecule by introducing amino acid residues (e.g., histidines) able to specifically bind with metal ions. Catalytic activity and stability of such metal-chelated enzymes in organic solvents is significantly higher than in the case of their non-modified analogs. 3.
INCREASE OF ENZYME ACTIVITY IN NON-AQUEOUS MEDIUM
A serious limitation on the way to a broader practical application of non-aqueous enzymology is in the low catalytic activity of enzymes in neat organic solvents as compared to water [3]. Because enzymes are otten insoluble in organic solvents, most commonly they are utilized as suspensions of lyophilized protein powder prepared by freezing aqueous enzyme solution in liquid nitrogen and drying m v a c u o . At different steps of this preparation, the enzyme is subjected to multiple stresses, which unavoidably decrease its catalytic potential [ 18]. During freezing, the enzyme can be cold-denatured at low temperature or due to contact with ice. Dehydration stress during drying leads to conformational changes, which are fastened by multiple inter-protein contacts in the powder and due to low protein mobility in the absence of water [ 19]. Finally, contact with anhydrous solvents may additionally dehydrate the enzyme by "stripping off" essential water [20]. An easy way to avoid some of these stresses is to use the methods other than freezing and vacuum drying. For example, enzyme solution can be deposited onto porous support and dried at normal pressure. High activities are obtained with cross-linked enzyme crystals suspended in organic solvents [9]. One more possibility is to apply organo-soluble complexes of enzymes with surfactants or lipids, whose activity in non-aqueous media is sometimes close to that in the aqueous solution [21]. Catalytic activity can also be increased when a traditional freeze-drying method is used and some examples of such an activation are discussed below.
3.1.
Engineering of protein matrix by co-lyophilization with excipients
Significant progress in enzyme activation in non-aqueous media has been achieved when enzymes have been lyophilized together with high- and low-molecular-weight compounds of different nature. Especially impressing activation was observed for subtilisin Carlsberg colyophilized with (,g21 [22]. The value of ke.at/gm for transeeterifieation reaction in hexane increases pronounoedly with increase in salt content and the activation reaches a factor of thousand in the powder containing 98% of the salt and 1% o,f the enzyme, Fig. 3 A. Such a significant activation has been studied in more detail and the following questions have been put forward: Is this activation effect general to other enzymes and excipients? How strongly is this effect influenced by solvent nature, protein hydration and other factors7 And generally, what mechanisms are behind this activation phenomenon? Thermolysin presents another example of an enzyme with a very polar transition state. Lyophilization with salt significantly activates this metallo-peptidase in the reactions of peptide synthesis in neat organic solvents - more than 200-fold activation is achieved in the powder containing 99% KCI and 0.5% of thermolysin. Hundred-to-thousand-fold activation by KCI of the two proteases, subtilisin and thermolysin, is not a result of trivial relaxation of substrate
360
4
A
2 iiJII- ----B~in-'~-il'-''~-gi / i/ / 4 i/B~UB 1 /
2~ %lql
!
4
I
,
,
0
2o
40
i
6o
I
8o
~.f i
100
0
~.
,,~n,,t~.
~
i
i
f
f
r
1
2
3
4
5
6
Fig. 3. Activation of proteases in organic solvents by lyophilization with KC1. A - dependence of the logarithm of the rate of subtilisin Carlsberg-catalyzed transesterification of N-benzoyl-Ltyrosine ethyl ester with propanol in hexane in matrices with KCI (Agc0, in relation to the reaction rate in protein powder (Ao) on the content of KCI in the matrices. B - dependence of the logarithm of the rate of peptide synthesis from furylacryloyl glycine and L-leucine amide catalyzed by thermolysin in tert-amyl alcohol on the content of water in the solvent. 1 - the powder with 99% of thermolysin and 1% of buffer components; 2 - the powder with 98% KCI, 1% ofthermolysin and 1% of buffer components. diffusional limitation in catalysis due to enzyme "dilution" by the salt - this has been shown in experiments on co-lyophilization of active enzymes with catalytically inactive proteins [23]. Subtilisin activity in neat organic solvents has appeared to be very sensitive to the nature of salts used as excipients. While all concentrations of salting-out anions such as fluoride, phosphate and acetate activate the enzyme, the activity is significantly suppressed by a salting-in thiocyanate. A good correlation has been observed between enzyme activation by the salt matrix and the anion position in the Hoffmeister series (Mozl~ev et al., submitted). Activation by salt is very sensitive to water content in organic solvents. In 98% KCI, the activation effect for thermolysin is equal to a factor of 100 in neat tert-AmOH, it increases to more than 2,000 on addition of 2.5 vol. % of water and then drops to less than 10 at more than 4 vol. % of water. This dependence has been explained by the fact that in the salt matrix thermolysin is significantly activated when the very first portions of water (up to 0.5 vol. %) are added (Fig. 3B). On the other hand, an especially fast increase in activity of the enzyme without salt is observed between 2.5 and 3 vol. % of water. Probably due to the charged nature of the salt matrix, high polarity in the enzyme environment necessary for high catalytic activity is already achieved at small water amounts, while much more water is needed to obtain equally high polarity in less polar protein powder. Oxidation of guaiacol by peroxidase is another reaction significantly accelerated by salt activation factor of 100 is obtained in 95% KCI. However, dependence of this activation on addition of water to acetonitrile is weaker, probably because the transition state of peroxidasecatalyzed free radical oxidation is much less polar than that in the proteases. Activation by salt depends on the organic solvent. The rate of transesterification by subtilisin in 95% KCI is zero in such a polar solvent as DMF but it increases significantly with an increase in the solvent's hydrophobicity. The activation by salt has a similar dependence on solvent log P: it is either absent or very small for polar solvents but reaches a factor of 100-to1000 in non-polar solvents. Similar dependence on solvent polarity has been obtained for the
361 activation of thermolysin. Most probably the highly polar salt matrix creates a favorable polar media for the enzyme reaction that produces significant enzyme activation in non-polar solvents. In contrast, because of the high polarity, the salt matrix may promote attraction of the molecules of polar solvents, thus provoking enzyme denaturation due to "stripping off" the essential water of the enzyme molecules (Mozhaev et al., submitted). Enzyme activation strongly depends on the nature of exeipients. By using tetrabutyl ammonium chloride, a lyodeproteetant, subtilisin has been activated although to a smaller extent than by KCI. On the other hand, almost all quantities of the lyoprotectant sucrose have depressed the activity of subtilisin in hexane and lyophilization with another lyoprotectant, BSA, has decreased activity of thermolysin. In combination, all these data indicate that the activation by exeipients does not correlate with their lyoprotectant efficiency and can not be explained by protection of enzyme against denaturation during the freezing and drying steps. Thousand-fold activation effects are obtained for the enzymes freeze-dried with PEG this excipient is almost as effective as KCI in activating subtilisin and thermolysin. Similar to KC1, activation by PEG strongly depends on solvent and water content indicating that the activation mechanisms of these two excipients may be similarly related to their polar character. 3.2.
Activation of enzymes by lyophilization with denaturants
Among different molecular reasons behind low activity of enzymes in non-aqueous media, one of the most important is a decreased molecular flexibility of the protein molecule in the absence of sufficient amount of water, which acts as a lubricant in macromolecular motion. One of the possibilities to intensify this motion can be achieved by reducing disulfide bonds, which cross-link many proteins and increase their molecular rigidity. To find out whether reduction of disulfide bridges may activate enzymes in non-aqueous solvents, a protease, c~chymotrypsin, containing 5 S-S bonds, has been lyophilized with different amounts of dithiothreitol (DTT) and its activity in transesterification reaction has been studied. In non-polar solvents ct-chymotrypsin freeze-dried with DTT shows the same activity as the enzyme in the absence of excipients, however 10-30-fold activation effects are obtained in polar solvents. A good correlation was observed between the activation effect and solvent hydrophobicity for a homologous series of alkyl acetates, C2-C6, used as solvents. Protein mobility is known to be decreased by polar solvents due to "stripping off" of water bound to the protein and this deleterious effect is probably compensated by additional flexibility achieved by reduction of disulfide bonds in the enzyme. The activation effect of DTT has been abolished by addition of water into organic solvents. This fact may be explained by the competition between water and DTT, which both create additional possibilities for protein dynamics, the latter being importantly involved in the enzyme catalysis. The activation shows a maximum at some intermediate concentration of DTT (10 raM) in the lyophilization buffer, while higher amounts of the thiol completely abolish the activity of ct-chymotrypsin in organic solvents. It is likely that only a limited number of S-S bonds is split at this optimal concentration of DTT (maybe the only one which supports the enzyme active site structure and is the most reactive among others), providing the enzyme with some optimal flexibility and activity. On the other hand, high concentrations of the thiol may completely destroy the unique three-dimensional conformation of the enzyme. Presence of free SH-group in the molecule of the excipient is essential for activation of ct-chymotrypsin: 10-fold enzyme activation is obtained by freeze-drying with 10 mM of other
362 thiols such as dithioerithritol and reduced glutathione. On the other hand, diols (ethylene glycol and butanediols), possible structural analogs of DTT without SH group, and oxidized glutathione do not show any positive effect on the activity of ~t-ehymotrypsin in organic solvents. In combination these facts indicate that activation of ot-ehymotrypsin in non-aqueous media is connected with S-S reducing ability of the thiols and is most likely explained by additional flexibility of protein conformation due to S-S bond reduction. If molecular flexibility is so important for enzyme activity/activation in non-aqueous media, then addition of small amounts of denaturants like urea and guanidinium chloride, which are known to loosen intra-protein contacts may activate enzymes as well. In fact, more than 10fold activation has been achieved for a-ehymotrypsin freeze-dried in 30 mM aqueous urea solution. The effect is decreased with an increase of the concentration of urea and at 1.5 M urea enzyme deactivation is observed. Another enzyme, subtilisin Carlsberg was activated 8-fold as a result of eo-lyophilization with moderate concentration of urea. This fact may indicate that activation of enzymes by additives that increase protein flexibility in anhydrous media is a rather general phenomenon. In conclusion, lyophilization of enzymes with exeipients may present a valuable approach to signitieantly enhancing catalytic activity of the enzyme catalyst. This earl be achieved either by engineering the favorable mieroenvironment of enzyme molecules or by stimulating molecular dynamics of the bioeatalyst through interactions with different effectors.
Acknowledgements The author is grateful to Drs. A.V. Levashov, A.K. Gladilin, E.V. Kudryashova, A.A. Vakurov (all Moscow State University), P.J. Hailing and D.G. Rees (University of Strathclyde, UK), J.S. Dordiek and his research group at the University of Iowa and Yu.L. Khmelnitsky (Enzymed Inc.) for collaboration in different parts of this research. This work was partially funded by grants from INTAS and US Army Research Office.
REFERENCES
.
3. 4. 5. .
7.
.
10. 11.
A.M.P. Koskinen and A.M. Klibanov (eds.), Enzymatic Reactions in Organic Media, Blaclde Academic & Professional, London et al., 1996. S.J. Singer, Adv. Prot. Chem. 17 (1962) 1. A.M.Klibanov, Trends Biotectmol. 15 (1997) 97. Y. Tomiuchi, T. Kijima and H. Kise, Bull. Chem. Soe. Jpn. 66 (1993) 1176. V.V. Mozhaev, Yu.L. Khmelnitsky, M.V. Sergeeva, A.B. Belova, N.L. Klyachko, AV. Levashov and K. Martinek, Eur. J. Biochem. 184 (1989) 597. J.A. Rupley, E. Gratton and G. Careri, Trends Biochem. Sci. 8 (1983) 18. Yu.L. Khmeluitsky, V.V. Mozhaev, A.B. Belova, M.V. Sergeeva and K. Martinek., Eur. J. Bioehem. 198 (1991) 31. V.V. Mozhaev, M.V. Sergeeva, A.B. Belova and Yu.L. Khmelnitsky, Biotechnol. Bioengin. 35 (1990)653. A.L. Margolin, Trends Biotechuol. 14 (1996) 223. A.K. Gladilin, E.V. Kudryashova, A.V. Vakurov, V.A. Izummdov, V.V. Mozhaev and A.V. Levashov, Biotechnol. Lett. 17 (1995) 1329. E.V. Kudryashova, A.K. Gladilin, A.V. Vakurov, F. Heitz, A.V. Levashov and V.V. Mozhaev, Bioteehnol. Bioengin. 55 (1997) 267.
363 12. 13. 14. 15. 16. 17. 18.
19. 20. 21. 22. 23.
Yu.L. Khmelnitsky, V.V. Mozhaev, A.B. Belova and A.V. Levashov, FEBS Lett. 284 (1991) 267. V.V. Mozhaev, E.V. Kudryashova, N.V. Efremova and I.N. Topchieva, Biotechnol. Teehniq. 10 (1996) 849. Kuehner and F.H. Arnold, Trends Biotechnol. 15 (1997) 523. C.-H. Wong, Trends Bioteehnol. 10 (1992) 378. F.H. Arnold and J.-H. Zhang, Trends Bioteehnol. 12 (1994) 189. R.D. Kidd, H.P. Yennawar, P. Sears, C.-H. Wong and G.K. Farber, J. Am. Chem. Soc. 118 (1996) 1645. J.F. Carpenter, S.J. Prestrelski, T.J. Anchordoguy and T. Arakawa, in: Formulation and Delivery of Proteins and Peptides, J.L. Cleland and R. Langer (eds.), ACS Symosium Series 567, Washington, pp. 134-147 (1994). K. Griebenow and A.M. Klibanov, Proc. Nat. Acad. Sci. USA 92 (1995) 10969. A. Zaks and A.M. Klibanov, J. Biol. Chem. 263 (1988) 8017. Y. Okahata and T. Moil, Trends Bioteehnol. 15 (1997) 50. Yu.L. Khmelnitsky, S.H. Welch, D.S. Clark and J.S. Dordick, J. Am. Chem. Soc. 116 (1994) 2647. B. Bedell, V.V. Mozhaev. D.S. Clark and J.S. Dordick, Biotechnol. Bioengin., in press.
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Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
365
Inactivation of enzymes at the aqueous-organic interface Peter J Hailing, Alistair C Ross and George Bell Departments of Bioscience & Biotechnology, Pure & Applied Chemistry and Chemical & Process Engineering, University of Strathclyde, Glasgow G1 lXW, UK p.j. halling@strath, ac. uk A liquid-liquid bubble column can be used to quantify interfacial inactivation of enzymes by organic liquids. Extent of inactivation per unit area can be greater for more hydrophobic solvents, particularly those with no polar groups. Inactivation rates can differ considerably even between similar enzymes, and do not show a maximum at the isoelectric point (unlike adsorption). 1. Introduction It is often desirable to use enzymes in the presence of an organic liquid phase (Klibanov, 1997; Carrea et al, 1995; Koskinen & Klibanov, 1995). One problem in such reaction systems may be inactivation of the enzyme by the organic species. To minimise inactivation in reactors, it is useful to understand the mechanisms involved. It is well established that 3 distinct mechanisms may contribute: 1) An indirect effect via removal of water from the microenvironment of the enzyme. This will not be further discussed here. 2) Interaction of individual organic molecules with the enzyme, leading to inhibition, conformational change, or other problems. 3) Interaction of the enzyme molecules with the organic phase interface, usually leading to major conformational change. Mechanisms 2 and 3 are most clearly distinguished in two-liquid systems, where there is a distinct aqueous phase (perhaps of very small volume). Such systems are useful for many applications, where at least one reactant or product has to be transferred via an aqueous phase (Freeman et al, 1993; Van Sonsbeek et al, 1993), or the enzyme is only active at high water levels. Mechanism 2 then corresponds to the effects of organic molecules dissolved in the aqueous phase which then interact with the enzyme. This is clearly distinct from direct contact of the enzyme with the phase interface (mechanism 3, interfacial inactivation). Experimentally, inactivation can be measured in systems of the same composition, but with different extents of agitation and hence interfacial area. It is often found that interfacial inactivation becomes the dominant mechanism in agitated systems, like typical reactors. However, interfacial inactivation remained poorly characterised, because the actual interracial area generated remains unknown in usual agitated reactors. We have developed a liquid-liquid bubble column apparatus that allows us to expose
366
enzyme solutions to known areas of interface under well defined flow conditions. This has allowed quantification of inactivation rates as a function of conditions, such as the type of solvent or enzyme. 2. The bubble column apparatus Fig 1 shows a schematic diagram of the apparatus used. Full details may be found in Ghatorae et al (1994a). Key features worth stressing are: 9 The use of hydrostatic head pressure to drive organic solvent through the bubble column, giving smooth flow. The pump is used only to re-cycle solvent. 9 The pre-column to ensure that the organic solvent is pre-saturated with water at the operating temperature. 9 The specially made nozzle to ensure small droplet sizes. 9 The coalescence device to ensure separation of clear organic solvent at the top of the column. In this apparatus, new interfacial area is continually created at the nozzle. After each droplet detaches, it will rise through the enzyme solution (over about ls). At the top of the column it normally persists for a similar period, before coalescing with the bulk organic liquid layer, with destruction of most of its interfacial area.
Fig. 1. The liquid-liquid bubble column apparatus
The column has an internal diameter of 7 mm, and the test solution is typically about 3 ml.
3. Inactivation depends on total interfacial area exposed, not time Experiments under given conditions are routinely repeated with different flow rates (by changing reservoir height). This changes the number of droplets passed through the solution per unit time, and the rate of exposure to interfacial area. Fig. 2 shows a typical example of the behaviour that has been found in every case studied. The enzyme activity declines more quickly as the droplet rate is increased. If the residual activity is plotted against the total interracial area to which the solution has been exposed, all points fall on the same line. Thus a fixed amount of enzyme will always be inactivated by a given area of interface, and the ratio between them
367
6-( E v
o m ~ .,...
6-{ (?,
5-
E 5-
.,J
4-
m 4-
v
3-
>" 3 -
. m
lj
. ,.,.,
13 2 -
0
"~ 2 -
t
0
I
i
2000 time (min)
I
4000
0 0
i 1
I 2
I 3
l 4
I 5
6
total surface exposed (m 2)
Fig. 2. Kinetics of inactivation of chymotrypsin at pH 6 by tridecane.
Droplets were passed at 25 (0), 64 (r-i) or 122 (V) min "1, or only initially for a solventsaturated control (z~). Ross et al, in preparation.
characterises the behaviour for the combination of enzyme, solvent and conditions. The measured gradient, with units of activity per unit area, can be taken as the rate of interfacial inactivation. Using the specific activity of the enzyme, the rate may be converted to mass of enzyme per unit area. Such rates are often found to be of the order of a few mg m "2, the density of a protein monolayer. These findings suggest that each droplet accumulates a more or less complete adsorbed layer of protein during its passage through the enzyme solution. This occurs independently of the dissolved protein concentration in the range we have studied (the zero order progress of inactivation is one piece of evidence here). Formation of an essentially complete monolayer under our conditions is consistent with the typical rates of protein adsorption measured in surface trough experiments (Dickinson et al, 1988), or with solid particles (Norde, 1995). All (or at least a constant fraction) of the protein in the adsorbed monolayer must now be destined to be inactivated. However, our experiments cannot show whether denaturation or inactivation occur immediately on adsorption, during the lifetime of the adsorbed layer, or when it collapses as the droplet coalesces at the top of the column. In general, when interfacial inactivation is measured, we also observe an accumulation of whitish, glutinous, insoluble particles. In addition, protein assays show that dissolved protein disappears from the aqueous solution at a similar rate to activity. Thus we can suggest that most of the inactivated enzyme has been converted to an insoluble aggregated form by the action of the interface. In some cases significant inactivation will still be found as droplet rates become very low or zero. This can be attributed to effects of dissolved organic solvent molecules (mechanism 2 in the introduction). Small rates of dissolved solvent
368
inactivation can be corrected for, before analysing interfacial inactivation. We generally try to avoid systems where dissolved solvent inactivation occurs at rates comparable with the interracial process. Protein inactivated by dissolved solvent may adsorb to the interface and protect other enzyme molecules, so correction for the dissolved solvent effect is uncertain. 4. Influence of solvent selection: more hydrophobic can be worse
Table 1 shows the rate of interracial inactivation of three enzymes by different organic solvents having a wide range of hydrophobicity. There is no simple trend for all enzymes. It is generally thought that biocatalyst stability is always highest with the least polar and/or most hydrophobic solvents. The octanol-water partition coefficient, log P, is the most commonly used parameter, and better stability with high log P has been observed in many studies. However, the interfacial inactivation of chymotrypsin shows clearly the opposite trend, with the most hydrophobic solvents being the most inactivating. Such opposite trends with log P had previously been noted in some studies of agitated systems (e.g. Cantarella et al, 1991 ). With urease there is no clear trend, while the lipase is most inactivated by the most polar solvents. We may rationalise these contradictory trends in terms of the mechanisms involved. Greater stability with high log P solvents is normal in cases where dissolved solvent mechanisms of inactivation predominate. The more hydrophobic the solvent, the lower the concentration present in the aqueous phase and seen by the biocatalyst. (There is a very close correlation between log P and saturated solubility in water.) Hence, it is reasonable to expect lower inactivation. A similar argument can be made in terms of partitioning of solvent molecules between the bulk organic phase and sites on the enzyme molecules. When inactivation takes place at the interface, however, the position is different. The adsorbed enzyme is now more or less equally exposed to the organic liquid, whatever its hydrophobicity. Under these circumstances the intrinsic inactivating tendency of the solvent will dominate, and it is reasonable to think that a less polar solvent will be more damaging. Table 1. Effect of solvent selection on interfacial inactivation
. Solvent 1-butanol isopropyl ether 2-octanone n-hexane n-butylbenzene n-tridecane
Interracial inactivation (pkat m"2)
log P
Urease
Lipase
Chymotrypsin
0.8 1.9 2.4 3.5 4.1 7.1
0.84 0.67 < 0.2 0.54 < 0.2 0.51
28.8 2.9 1.7 < 0.6 < 0.6 < 0.6
0.95 < 0.5 < 0.5 4.95 3.9 5.1
Jack bean urease, Candida rugosa lipase, and bovine (z-chymotrypsin. Ghatorae et al, 1994b.
369
The opposite trend for interfacial inactivation of the lipase probably reflects evolutionary selection for adsorption and stability at the substrate-water interface. The triacylglycerol substrate of lipases is relatively non-polar, so we would not expect rapid inactivation at hydrophobic interfaces.
5. Solvent selection: amphiphilic molecules better Table 2 shows the rates of interracial inactivation of chymotrypsin by a selection of solvents with similar log P values, but differing in the functional groups contained. As can be seen, their inactivating tendencies are significantly different. In particular, inactivation is much less for the strongly amphiphilic solvents decanol and undecanenitrile. These have a terminal very polar functional group, balanced by a longer hydrocarbon chain to give the same log P. The branched and secondary alcohol 3,7-dimethyl-3-octanol is less amphiphilic and more inactivating. This makes sense when we consider that molecules with an amphiphilic character will tend to be strongly oriented at the interface, with the polar groups directed towards the aqueous phase. Thus the enzyme will see an interface made up largely of polar groups, and will have little contact with the hydrophobic parts of the solvent molecules. A related feature of such interfaces will be a lower interracial tension, as seen from the values in Table 2. A correlation with interfacial tension for inactivation by organic solvents in agitated systems was suggested by Owusu & Cowan (1989).
Table 2. Interracial inactivation of chymotrypsin by different solvents of similar hydrophobicity. Solvent heptane 1-octene cyclooctane butyl benzene 1-chlorohepta ne isoamyl ether ethyl octanoate 2-undecanone 3,7-d imethyl-3-octanol undecanenitrile decanol
Interracial inactivation (mg m2)
Interfacial tension (mN m~)
0.78 0.84 0.64 0.60 0.78 0.56 0.39 0.74 0.34 0o16 0.14
50.2 50 50 41.4 35 32.2 25.5 23.9 20 19 10.2
All these solvents were selected as having log P between 3.8 and 4.1. Undecylenic acid gave a much higher rate of inactivation (1.53 mg m2), which may reflect a specific effect of the COOH groups. Ross et al, in preparation.
370 Amphiphilic molecules will tend to accumulate at the interface even when they are only one of the components of the organic phase (i.e. they will be surfactants). When a 50% mixture of decanol and heptane was used under the conditions of Table 2, the rate of inactivation was 0.23 mg m2, little more than with pure decanol. Such mixed solvents may be particularly useful to control interfacial phenomena. Feliu et al (1995) have described prevention of enzyme inactivation in an agitated system by added surfactants. A series of experiments with the related enzyme 13-chymotrypsin showed similar behaviour, with inactivation rates falling from 1.00 mg m"2 with heptane to 0.31 mg m"2 with decanol. With a third enzyme, pig liver esterase, the trend was much less clear, though decanol again gave the lowest inactivation rate of five solvents studied. 6. Inactivation is not maximal when pH = pl, unlike adsorption A maximum is usually found in interfacial adsorption of a protein when the pH is equal to its isoelectric point (Norde, 1995; Dickinson et al, 1988). Away from the pl, the net charge of the adsorbed protein will tend to oppose further adsorption of protein molecules from solution, as these have the same sign charge. Fig. 3 shows the interfacial inactivation of three serine proteases as a function of pH. The enzymes are closely related, but have very different pl values. It is clear that pH has a large influence on the rates of inactivation. However, it is equally clear that inactivation does not show an obvious maximum at the pl. This is probably because inactivation depends on the extent of structural change at the interface, as well as on the amount of protein adsorbed. There is some 1.o evidence that the conformational changes may often be less near the pl, in .> part because of the closer packing of the c adsorbed molecules (Kondo et al, 1992). ~ 0.5 There are probably also contributions from the effects of pH on the general ID pl ~-CT pl e~-CT stability of the native fold. There may be pl "[ some tendency to a maximum in , 1 . . . . I , , 1 0.0 inactivation at pH around (pl + 1), and a 4 6 8 10 difference in effective pi on the surface pH might be a factor here. Interestingly, some measurements of Fig. 3. Rate of inactivation of serine foamability of these protein solutions as proteases as a function of pH. a function of pH have shown a rather o~-chymotrypsin (O), 13-chymotrypsin (rq) good correlation with effects on and trypsin (A). Acetate, glutamate, Tris interfacial inactivation (Ross et al, in and imidazole buffers were used. Ross et preparation). al, in preparation.
E1.5
~
04
371
7. Critical enzyme properties are unclear Table 3 summarises the rate of interfacial inactivation by n-alkanes of all the enzymes we have studied so far. Some enzymes are clearly more prone to interfacial inactivation than others, but the critical protein properties are not obvious. Ribonuclease and papain stand out as relatively resistant, and compared with most of the other proteins studied they also have higher temperatures for thermal denaturation and lower hydrophobicity. Low values of partial specific adiabatic compressibility have often been related to less unfolding at interfaces ("hard" versus "soft" proteins; Kondo et al, 1992; Norde, 1995). However, the correlation with our inactivation data is not strong, whether using experimental or predicted compressibilities. One problem in looking for correlations is the effect of other conditions. For example, we have shown above that interfacial inactivation can be a strong function of pH. This can lead to very different conclusions about relative stabilities, e.g. ~chymotrypsin is inactivated at only one third of the rate of 13-chymotrypsin at pH 5.0, but more than 2 times faster at pH 9.0. Many enzyme properties are also greatly affected by pH, and this includes thermal denaturation and compressibility (as recently shown by Chalikian et al, 1995).
Table 3. Rate of interfacial inactivation of different enzymes by n-alkanes. Enzyme
cx-chymotrypsin 13-chymotrypsin trypsin urease pig liver esterase ribonuclease papain
Interfacial inactivation by alkanes (mg m"2) 0.78-1.00 0.94-1.00 0.50 1.00 0.62 <0.15 <0.20
Hydrophobicity
Compressibility (10 "12 cm2dyn"~)
44 42 46 40-60
0.131 0.169 0.122 0.06 0.165
4.15 7.0* 0.92 -9.3* 6.5*
62 66
-0.13 0.023
1.12 -4.6*
Molecular Tm (~ mass (kDa) 25.2 22.5 23.4 546 165 13.7 23.4
Rates shown are from experiments with either hexane, heptane or tridecane; these solvents have been found to give similar rates of inactivation for any given enzyme and conditions. Lipase is excluded from the list because of its design for stability at such interfaces. Ross et al, in preparation. * calculated from the correlation of Gromiha & Ponnuswamy (1993) 8. Acknowledgment We thank BBSRC for the award of a studentship.
372
9. References
M Cantarella, L Cantarella, F Alfani (1991) Enzyme Microb. Technol. 13, 547-553 "Hydrolytic reactions in two-phase systems. Effect of water-immiscible organic solvents on stability and activity of acid phosphatase, beta-galactosidase and beta-fructofuranosidase" G Carrea, G Ottolina, S Riva (1995) Trends Biotechnol. 13, 63-70 "Role of solvents in the control of enzyme selectivity in organic media" T V Chalikian, A P Sarvazyan, K J Breslauer (1994) Biophys. Chem. 51, 89-109 "Hydration and partial compressibility of biological compounds" E Dickinson, B S Murray, G Stainsby (1988) In: E Dickinson, G Stainsby, Eds, Advances in Food Emulsions and Foams, Elsevier Applied Science, London, pp 123-162 "Protein adsorption at air-water and oil-water interfaces" J A Feliu, C Demas, J Lopez-Santin (1995) Enzyme Microb. Technol. 17, 882-887 "Studies on papain action in the synthesis of Gly-Phe in 2-liquid-phase media" A Freeman, J M Woodley, M D Lilly (1993) Bio/Technol 11, 1007-1012 "ln-situ product removal as a tool for bioprocessing" A S Ghatorae, G Bell, P J Hailing (1994a) Biotechnol. Bioeng., 43, 331-336 "Inactivation of enzymes by organic solvents: new technique with well-defined interfacial area" A S Ghatorae, M J Guerra, G Bell, P J Hailing (1994b) Biotechnol. Bioeng., 44, 1355-1361 "Immiscible organic solvent inactivation of urease, chymotrypsin, lipase and ribonuclease. Separation of dissolved solvent and interfacial effects" M M Gromiha, P K Ponnuswamy (1993) J. Theor. Biol. 165, 87-100 "Relationship between amino acid properties and protein compressibility" A M Klibanov (1997) Trends Biotechnol. 15, 97-101 "Why are enzymes less active in organic solvents than in water?" A Kondo, F Murakami, K Higashitani (1992) Biotechnol. Bioeng. 40, 889-894 "Circular dichroism studies on conformational changes in protein molecules on adsorption on ultrafine polystyrene particles" A Koskinen, A M Klibanov (1995) Enzymatic reactions in organic media, Chapman & Hall, Andover. W Norde (1995) Cells Materials, 5, 97-112 "Adsorption of proteins at solid-liquid interfaces" R K Owusu, D A Cowan (1989) Enzyme Microb. Technol. 11,568-574 "Correlation between microbial protein thermostability and resistance to denaturation in aqueous:organic solvent two-phase systems" H M Van Sonsbeek, H H Beeftink, J Tramper (1993) Enzyme Microb. Technol. 15, 722-729 "2-liquid-phase bioreactors"
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
373
Exploiting hysteresis for high activity enzymes in organic media
Johann Partridge, Peter. J. Hailing and Barry D. Moore Department of Pure & Applied Chemistry, University of Strathclyde, Thomas Graham Building, 295 Cathedral Street, Glasgow G1 I XL, Scotland, U.K.
Common methods for removing water from protein preparations such as freeze drying and air drying are shown here to be much more detrimental to enzyme activity than simple rapid dehydration. In a comparison of drying methods for cross-linked enzyme crystals of subtilisin Carlsberg, the highest rates of reaction in organic media were obtained with crystals dried by washing with anhydrous polar solvent. Crystals dried over molecular sieves in air or solvent showed significantly lower activity. In all cases, full activity could be recovered in aqueous buffer. When different solvents were used to dry the enzyme crystals, the catalytic rate was found to vary significantly- longer chain alcohols gave the highest rates while smaller "water-like" solvents such as methanol gave much lower rates. The study was extended to immobilised forms of subtilisin Carlsberg and ot-chymotrypsin. When these preparations were dried by solvent washing, high catalytic rates similar to those of crystals prepared by the same procedure were obtained. Catalytic efficiency in polar solvents was 1000 fold greater than that of freeze-dried powders. As with the crystals, subsequent drying over molecular sieves resulted in a substantial loss in enzymatic activity. The activities of solvent washed immobilised enzymes and of cross-linked crystals were found to vary significantly as a function of system water content.
1. Introduction In recent years it has been shown that enzymes exhibit novel properties in low water organic media, such as catalysis of reactions impossible in water, altered substrate specificity and increased thermostability [1]. This has led to an increased demand for enzyme preparations which exhibit high catalytic activity and reproducible selectivity [2-5]. The commonly used freeze-dried powders are known to exhibit low activity [6,7] but are often used because they can be obtained in one step from commercially supplied enzyme. Enzymes immobilised onto supports often show higher activity in organic media but the types of support material reported may not always be readily available and well controlled immobilisation procedures are often required [8-12]. Recently commercial cross-linked enzyme crystals (CLECs) have been introduced. As with enzymes immobilised on supports, these offer advantages over the conventional freeze dried preparations: access to individual enzyme molecules will be improved, whilst particle aggregation and diffusional limitation are considerably reduced. In addition, CLECs have been shown to exhibit high activity, good
374 stability and excellent reproducibility [13]. These characteristics make them potentially very attractive as catalysts for use in organic reaction mixtures. The method by which the biocatalyst is prepared for use in organic media often varies. With freeze dried powders, the enzyme which is essentially dry after lyophilisation, is added directly to the organic solvent which contains a known amount of water [14-16]. Similarly, enzymes immobilised from aqueous solution are collected by filtration, dried under vacuum and used directly in solvent with fixed water content. In other studies, the biocatalyst and solvent are dried exhaustively over water absorbents such as molecular sieves and then pre-equilibrated to well defined thermodynamic water activity (a,,), either together or separately prior to reaction [17,18]. This method offers advantages. Catalytic activity of a particular type of preparation is governed by the amount of water bound to the enzyme molecules, which is only a fraction of the total present. At equilibrium there will be a characteristic relationship between the amount of water bound and that dissolved in the organic phase. If the water level is expressed in terms of its aw, the relationship between catalytic activity and residual water level is generally similar for many different solvents [ 17,18]. There have been some literature reports that different catalytic activity can be found in systems with the same water content or a,,, but different hydration histories [19]. In this work, we set out to examine how different methods of enzyme pre-treatment effected their subsequent catalytic efficiency in organic solvent. Such studies on freeze dried enzymes are problematic. Random but significant differences in morphology between batches of powders is often found, probably due to variations in temperature and pressure during lyophilisation. Differences in catalytic rates can result, so that it can be difficult to make direct rate comparisons between laboratories and also to carry out meaningful studies of possible hysteresis effects arising from different treatments. We anticipated that using more homogeneous enzyme preparations these problems would be reduced. 2. Results and Discussion 2.1
The effect of different CLEC pre-treatments on catalytic efficiency Subtilisin Carlsberg CLECs (ChiroCLEC-BL) were supplied as a suspension in aqueous buffer. The manufacturer's literature reports that this suspension can be stored indefinitely at 4~ without any significant loss in catalytic activity. We were interested in studying the effect of different methods of preparing the crystals for use in organic media on their subsequent catalytic activity. In the present study, we routinely started with the fully active preparation in aqueous suspension and treated the samples by different methods for use in solvent as required. A number of routes can be envisaged for moving from an aqueous suspension to a "dry state". Whilst lyophilisation is necessary when drying dissolved enzymes for use in organic solvents, it is not the simplest and most obvious choice of water removal when enzymes are used as CLECs. Besides this, one of the main reasons for the low activity exhibited by freeze dried powders in organic media is known to be the deleterious effects of dehydration by this method. This process is therefore best avoided. Water can be removed from CLECs by more simple methods, including washing with anhydrous polar solvent and drying over molecular sieves in air or in solvent. As a starting point we explored the effect of washing CLEC with either propanol or acetonitrile. The solvent washed enzyme was then used directly in a reaction, or preequilibrated over molecular sieves in air or solvent (methods commonly applied to
375 conventional freeze dried and immobilised enzymes). We followed the rate of a standard transesterification in anhydrous acetonitrile using CLECs pre-treated in different ways: the initial reaction rates are shown in Table 1. The highest rates were obtained with CLEC placed directly in the reaction mixture atter solvent washing, whilst equilibration at fixed water activity invariably reduced the rate. Although CLEC pre-equilibrated in air gave by far the lowest rate, this preparation is still more active than the conventional freeze dried preparation of subtilisin (<0.01nmol/mg/min). Interestingly, CLEC pre-equilibrated in anhydrous acetonitrile over molecular sieves gave higher transesterification rates. Such an observation has been previously reported in the literature [21]. Freeze dried subtilisin powder in hexane was shown to give higher rates when equilibrated in solvent as opposed to in air. However, the difference between the two treatments is much greater for the CLECs in acetonitrile. Table 1. Effect of enzyme treatment on catalytic activity of subtilisin Carlsberg CLEC in anhydrous acetonitrile. Washin B Solvent" Further Enzyme Treatment ~ PrOH none PrOH 1 PrOH 2 ACN none ACN 1 ACN 2 PrOH b none PrOH, AcN none AcN, PrOH none AcN, aqueous buffe,r, PrOH .. none
Rate (nmol/m~min) 224 • 7 34 + 4 2.9 + 0.1 112 + 4 33 + 3 2.3 + 0.2 187 + 2 84 • 4 102 + 3 190 + 14
Data from Partridge et al [20] " CLEC was rinsed 3 times with 1 ml of each solvent shown respectively (with the exception of b which was rinsed 6 times). Atter each wash, the solvent was removed by centrifugation. c (1) Solvent washed CLEC was suspended in anhydrous AcN and then equilibrated over molecular sieves for 3 days, 20~ (2) Solvent washed CLEC was equilibrated over molecular sieves for 3 days, 20~ After this time it was suspended in anhydrous AcN. From Table 1 it is apparent that the difference in catalytic rate for CLEC dried over molecular sieves through the vapour phase is small, regardless of the solvent used to wash the crystals prior to equilibration. This is also the case for CLEC equilibrated in the reaction solvent prior to use. However, it is interesting to note that CLEC washed in propanol and used directly in the reaction gives a rate approximately double that of CLEC which was washed in acetonitrile. When the crystals were washed consecutively with propanol (x3) and acetonitrile (x3), regardless of the order in which the solvents were used, the rates of catalysis were less than half that for CLEC washed solely in propanol, but were close to that of acetonitrile washed CLEC. When the number of washes is increased from 3 to 6 for propanol washed CLEC, the decrease in rate is barely significant. It is apparent that the effect of acetonitrile is dominant on the crystals and the catalytic efficiency of an acetonitrile washed
376 CLEC can not be increased by subsequent washing with propanol. To ascertain whether or not acetonitrile washing had a long term detrimental effect on the crystals, we measured the catalytic activity of CLEC which was rinsed with acetonitrile, buffer and propanol respectively. The majority of catalytic activity was recovered, indicating that acetonitrile does not appear to damage the CLEC irreversibly. It might also be inferred that pre-equilibration through the vapour phase results in structural damage to the enzyme crystals, so that most of their catalytic activity is irreversibly lost. We attempted to eliminate this possibility experimentally. Biocatalyst which had been solvent washed and leit to equilibrate through the vapour phase for 3 days, was returned to an aqueous environment. After this, it was solvent washed to remove excess water, and used immediately to catalyse a reaction in anhydrous acetonitrile. CLEC was found to regain high catalytic activity. Furthermore, the rate observed is comparable to that of CLEC which has undergone solvent washing only. It is interesting to note that the catalytic efficiency of CLECs extensively dried over molecular sieves can not be improved significantly by further equilibration over a high aw saturated salt. CLEC must be returned to the aqueous buffer if high catalytic activity is to be recovered. The observations discussed above are not exclusive to reactions in acetonitrile. Experiments in propanol have confirmed that enzyme which has undergone solvent washing has a catalytic rate more than 12-fold higher than that which has been dried further using molecular sieves. Similarly, for reactions in propanol, enzyme pre-equilibrated through the vapour phase can regain high catalytic activity by exposure to an aqueous environment. The results discussed above clearly demonstrate that the catalytic behaviour of CLECs exhibits pronounced hysteresis: rates vary by 80-fold depending on previous hydration history. One might speculate that the enzyme or enzyme crystal has a 'memory' of how it has been treated, and this 'memory' can only be erased by returning the CLEC to an aqueous environment. Two possible explanations may account for this observation: (1) different dehydration protocols effect the amount of water left bound to the enzyme, or (2) the conformation of the dried enzyme is very sensitive to the method of water removal. Most probably these two effects are intimately related. Since washing the CLEC with solvent gives the most efficient rates of catalysis, we went on to investigate the effect of washing with other solvents. Table 2 shows how transesterification rate in anhydrous acetonitrile varies as much as 7-fold depending on the organic solvent used to dry the catalyst. The alcohols generally gave the fastest rates of catalysis, followed by acetonitrile and acetone which gave intermediate rates. Interestingly, the smaller "water-like" solvents, ethane diol and methanol gave the lowest rates of all solvents tested. With CLEC washed in methanol, only 33% of the maximum activity is regained after brief washing with buffer and propanol. Leaving in buffer overnight resulted in increased catalytic activity (63% of the original propanol washed CLEC). Nevertheless, the catalytic performance of all solvent washed CLECs was still higher than that which had been equilibrated in air or in solvent at aw < 0.01. A plausible explanation for this variation in catalytic rate when different solvents are used to dry the CLEC is that each solvent has a very specific capacity to displace the water within the crystal. Conformational changes may result on removal of more water from the crystal leading to a less active enzyme state and decreased rates of catalysis. On the basis of these data one can conclude that care must be taken when choosing a solvent to wash the crystals if maximum enzymatic activity is to be achieved.
377 Table 2. Effect of different solvent washes on the catalytic activity of subtilisin Carlsberg CLE C in anhydrous acetonitrile.
Washin~ Solvent' EtOH BuOH PrOH Me2CO ACN Ethane Diol CH3 OH MeOH, aqueous buffer ,. PrOH
Rate (nmol/mg/min) 282 + 32 243 • 23 224 • 7 121 + 11 112 + 4 50 + 12 39+ 3 ...... 74 + 1
Data from Partridge et al [20] ' CLEC was rinsed 3 times with 1 ml of each solvent shown respectively. After each wash, the solvent was removed by centrifugation. The enzyme was then suspended in anhydrous acetonitrile. 2.2
Extension of our findings to immobilised preparations With dissolved enzymes water must generally be removed by freeze-drying or similar prolonged dehydration methods. With CLECs, rapid drying by solvent rinsing is possible and this appears to lead to better specific activity. We therefore hypothesised that if non-crystalline but immobilised enzymes were treated using the same solvent washing procedure, very high activities might be obtained. We tested this hypothesis for an immobilised form of subtilisin Carlsberg. The enzyme was prepared using a simple adsorption procedure from aqueous buffer (pH 7.8) unto a standard silica gel support. Table 3 shows a comparison of the initial reaction rates obtained in acetonitrile using subtilisin Carlsberg prepared in different forms. The propanol rinsed immobilised enzyme preparation (termed PREP) gave fairly low rates in anhydrous acetonitrile. Table 3. Effect of preparation type on subtilisin Carlsber~; activity in acetonitrile. Rate (nmol / nag "/rain) 0% H20 v/v in ACN 1% H20 v/v in ACN Enzyme form and treatment (aw < 0.01) (a, = 0.22)
freeze dried powder, air dried b CLEC, PrOH washed CLEC, PrOH washed, air dried b immobilised form, PrOH washed immobilised form, PrOH washed, air drie db
< 0.01 226 2.94 0.82 ....
0.13 610 13.8 142 0.60
Data from Partridge et al [20,22] ' refers to weight of enzyme in preparation, b sample placed in sealed jar over molecular sieves for 3 days to give enzyme at aw<0.01; further 3 days equilibration over H20-saturated potassium acetate was carried out for enzyme at a, = 0.22.
378 However, on addition of 1% water v/v to the reaction solvent, the PREP was found to exhibit high activity. At this water level, rates for the PREP were found to be comparable to the CLEC, and over 1000 times greater than the commonly used freeze-dried powders. If PREP was subsequently dried in air prior to assaying in the organic solvent most of the activity was lost and the residual level approached that obtained for the lyophilised powder (see Table 3). However, aqueous suspensions of the silica adsorbed enzyme can be stored at 4~ for at least 3 weeks with negligible loss of activity and converted to the PREP as required. The effect of washing the immobilised enzyme with different anhydrous organic solvents was also tested. The same pattern of results as that observed for the CLEC emerged: ethanol washing gave high rates, acetonitrile gave intermediate rates and methanol produced a much less active preparation. Cross-linked enzyme crystals of chymotrypsin are not commercially available but under the same reaction conditions described in Table 3 the PREP of this enzyme exhibited high catalytic activity in acetonitrile with water levels of 1%v/v and above. Again these rates were two orders of magnitude better than the freeze-dried powder. It is perhaps surprising the method described here for preparing conventional biocatalysts for reactions in low water media has not been reported previously. However, until recently the large hydration hysteresis effects obtainable with enzymes had not been fully recognised. Our work with CLECs [20] and immobilised enzymes [22] has shown that different methods of water removal can dramatically affect the enzyme activity obtained. 2.3
The effect of system water content on catalytic activity Previous studies with biocatalysts in organic media have shown that the amount of water present in the system plays an important role in controlling factors such as rate, stability and hydrolytic equilibria [ 1, 23]. We therefore proceeded to carry out a more detailed study of the variation in catalytic activity as a function of water level in the actual reaction mixture for subtilisin CLEC and the immobilised form which had been propanol washed. Figure 1 shows the rate profiles for propanol washed CLEC and PREP in acetonitrile as a function of the thermodynamic water activity, aw. The profile for the freeze dried form of this enzyme is also shown for comparison. However, it should be noted that since the activity per unit weight of enzyme is much lower in the lyophilised powder, the rate profiles for the three preparations are shown as relative rates normalised to their maximum values. According to previous studies the amount of water bound to a protein in solvent would be expected to be controlled by a~ [23]. Under these conditions differences in the amount of residual bound water should be eliminated and hence similar rate vs aw profiles might be expected for all three forms of the enzyme. As can be seen in Figure 1 this is not the case. The activity of the lyophilised powder continues to increase even up to aw 0.76, while a maximal rate is obtained at aw of 0.11 with the CLEC and aw of 0.44 with the PREP. The different rate profiles could arise because the three preparations of the enzyme differ in either water binding or the water required for catalytic activity. A large change in water binding isotherms is unlikely, but kinetic factors may be significant. The water content of the lyophilised powder will be determined by its adsorption isotherm. In contrast, treatment of the CLEC and PREP imposes a water desorption process, for which the isotherm will be different due to hysteresis, and even the apparent equilibrium value may not be reached in the time employed. An alternative hypothesis is that CLEC and PREP require less water because the propanol dehydration process leaves a large proportion of the enzyme molecules in
379 a conformation close to the active fOrm. Only low levels of water may then be needed to promote catalysis and further increases in water availability provide no beneficial effect. With the freeze-dried powder the very low rates suggest most of the enzyme is initially inactive. In this case a water catalysed reorganisation process is probably required to convert the enzyme back to an active state since much greater water levels are needed to obtain high activity. The difference in profiles in Figure 1 would therefore reflect the fact that water plays different roles in promoting enzyme catalysis depending on the hydration history of the system.
1.0 .
0.8
~> 0.6 "~
0.4 0.2 0.0 0.0
0.2
0.4
0.6
0.8
water activity, a,
Figure 1. Relative rate as a function of aw for the transesterification reaction in acetonitrile catalysed by propanol washed subtilisin CLEC (&), subtilisin PREP (r-3), and freeze dried subtilisin (O). Rates for each preparation were normalised relative to the maximum value: 610 nmol/mg/min at a~ of 0.11 for the CLEC, 159.7 nmol/mg/min at aw of 0.44 for the PREP and 3.3 nmol/mg/min aw of 0.76 for the freeze dried powder. Data from Partridge et al [20,22]. 3. Conclusions We have demonstrated that the catalytic behaviour of subtilisin CLECs in polar solvents exhibits pronounced hysteresis. The activity of solvent washed crystals was as much as 80-fold higher than that of crystals dried in air or solvent over molecular sieves. The solvent washing procedure was exploited to obtain high activity immobilised enzyme preparations of subtilisin Carlsberg and ct-chymotrypsin, enzymes with very different secondary and tertiary protein structures. This suggests the procedure may find widespread application as a simple and economical way of preparing biocatalysts for reactions in organic media. 4. Acknowledgement We thank J. J. Lalonde for helpful discussion. We are grateful to the Biotechnology and Biological Science Research Council for financial support.
380 References
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3. .
5. .
7. 8.
10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23.
A. Koskinen and A. M. Klibanov (eds), Enzymatic Reactions in Organic Media, Chapman and Hall, Andover, 1995. C. R. Wescott and A. M. Klibanov, Biochim. Biophys. Acta 1206 (1994) 1. J. Broos, J. F. J. Engbersen, I. K. Sakodinskaya, W. Verboom and D. N. Reinhoudt, J. Chem. Sot., Perkin Trans. 1 (1995) 2899. J. O. Rich and J. S. Dordick, J. Am. Chem. Soc. 119 (1997) 3245. G. A. Hutcheon, M. C. Parker, A. James and B. D. Moore, Chem. Commun. (1997) 931. K. Dabulis and A. M. Klibanov, Biotechnol. Bioeng. 41 (1993) 566. O. Almarsson and A. M. Klibanov, Biotechnol. Bioeng. 49 (1996) 87. A. O. Triantafyllou, D. B. Wang, E. Wehtje and P. Adlercreutz, Biocatal. Biotransfor. 15 (1997) 185. Z. Yang, A. J. Mesiano, S. Venkatasubramanian, S. H. Gross, J. M. Harris and A. J. Russell, J. Am. Chem. Soc. 117 (1995) 4843 E. Wehtje, P. Adlercreutz and B. Mattiasson, Biotechnol. Bioeng. 41 (1993) 171. J. F. Diaz and K. J. Balkus, J. Mol. Catal. B: Enzymatic 2 (1996) 115. M. T. Reetz, A. Zonta and J. Simpelkamp, Biotechnol. Bioeng. 49 (1996) 527. Y. F. Wang, K. Yakovlevsky and A. L. Margolin, Tetrahedron Lett. 37 (1996) 5317. A. Zaks and A. M. Klibanov, J. Biol. Chem. 263 (1988) 3194 A. Zaks and A. M. Klibanov, J. Biol. Chem. 263 (1988) 8017 S. H. M. van Erp, E. O. Kamenskaya and Y. L. Khmelnitsky, Eur. J. Biochem. 202 (1991) 379. R. H. Valivety, P. J. Hailing and A. R. Macrae, Biochim. Biophys. Acta 1118 (1992) 218 I. Svensson, E. Wehtje, P. Adlercreutz and B. Mattiasson, Biotechnol. Bioeng. 44 (1994) 549 P. J. Hailing, Enzyme Microb. Technol. 16 (1994) 178 J. Partridge, G. A. Hutcheon, B. D. Moore and P. J. Hailing, J. Am. Chem. Soc. 118 (1996) 12873 M. C. Parker, B. D. Moore and A. J. Blacker, Biocatalysis, 10 (1994) 269 J. Partridge, P. J. HaUing and B. D. Moore, Chem Commun. (1998) 841 G. Bell, P. J. Hailing, B. D. Moore, J. Partridge and D.G. Rees TIBTECH 13 (1995) 468.
Stability and Stabilization of Biocatalysts A. Ballesteros,F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998Elsevier Science B.V. All rights reserved.
381
L I M I T E D P R O T E O L Y S I S OF P R O T E I N S BY T H E R M O L Y S I N IN T R I F L U O R O E T H A N O L Patrizia Polverino de Laureto, Elena Scaramella, Marcello Zambonin, Vincenzo De Filippis and Angelo Fontana CRIBI Biotechnology Centre, University of Padua, Via Trieste 75, 35121 Padua, Italy
We have examined the proteolysis of model proteins by thermolysin when dissolved in aqueous buffer at neutral pH in the presence of 50% (by vol.) trifluoroethanol (TFE). Under these solvent conditions, proteins acquire a new conformational state characterized by enhanced helical secondary structure, but lacking the specific tertiary interactions of the native species. It was found that the TFE-state of proteins dictates very selective peptide bond fissions by the TFE-resistant thermolysin, which otherwise shows broad substrate specificity. Nicked protein species with a single peptide bond hydrolyzed have been prepared and isolated to homogeneity in the ease of bovine ribonuclease A (cleavage at Asn34-Leu35), hen lysozyme (Lys97-Ile98), bovine o~-lactalbumin (Ala40-Ile41) and horse cytochrome c (Gly56-Ile57).
1. INTRODUCTION Non aqueous enzymology has emerged as a very active and useful area of research in recent years, prompting fundamental questions concerning enzyme structure and dynamics and their effect on catalysis (Wong, 1989; Klibanov, 1989, 1997; Dordick, 1989; Mattiasson & Adlercreutz, 1991; Gupta, 1992; Carrea et al., 1995). The addition of water-soluble cosolvents may change dielectric constant, hydrophobicity and hydrogen bonding of the reaction medimn, altering the various forces responsible for the substrate binding specificity (electrostatic and van der Waal's forces, hydrophobic and steric effects, hydrogen bonds) (Jencks, 1969; Fersht, 1985; Kraut, 1988) and thus the catalytic properties of the enzyme. In particular, the hydrogen bonding properties of alcohols, sugars and polyols are expected to change both the solvent medium and the solvation of protein molecules, thus leading to (subtle) conformational changes of an enzyme and its catalytic behaviour (Pourplanche et al., 1994; Lign6 et al., 1997). In the case ofproteolytic enzymes, the cosolvent can alter the conformational properties of Abbreviations: CD, circular diehroism, SDS, sodium dodecyl sulfate, PAGE, polyacrylamide gel electrophoresis, TFE, trifluoroethanol, HPLC, high performance liquid chromatography, [0], mean residue ellipticity, nicked protein, a protein species with a peptide bond hydrolyzed.
382 both the enzyme and the peptide/protein substrate and consequently their mutual interaction. The use of proteolyfic enzymes in the presence of organic solvents has been extensively investigated in the past and successfully utilized for the synthesis or semisynthesis of peptides and proteins, since protease, in the presence of water-soluble cosolvents (glycerol, 2-propanol) catalyze the reverse reaction, i.e. the synthesis instead of hydrolysis of peptide bonds (Chaiken, 1981; Kullman, 1987; Wayne & Fruton, 1983; De Filippis & Fontana, 1990). Short-chain alcohols such as methanol, ethanol, propanol, chloroethanol and, in particular, tdfluoroethanol (TFE) have been shown to induce and stabilize tx-helical structure in otherwise randomly coiled or partially structured polypeptides (Tamburro et al., 1968; Nelson & Kallenbach, 1986; Lehnnan et al., 1990; Storrs et al., 1992; Stnnichsen et al., 1992). TFE is nowadays the cosolvent of choice for enhancing the helical secondary structure of polypeptides (Smith et al., 1994; Cammers-Goodwin et al., 1996; Bolin et al., 1996; Luo & Baldwin, 1997; Myers et al., 1998). The helixinducing effect of TFE does not seem to occur independently of the amino acid sequence of the polypeptide chain, since peptides and protein fragments corresponding to helical regions in the native protein have a higher tendency to form a helix in the presence of TFE (Lehnnan et al., 1990; Segawa et al., 1991). However, also protein fragments derived from predominantly J3-sheet proteins acquire a helical structure in the presence of TFE (Fan et al., 1993; Hamada et al., 1995; Jayaraman et al., 1996). TFE has been shown also to disrupt the native conformation of globular proteins (Shiraki et al., 1995, and references cited therein), but the resulting denaturated state is much different from the random-coiled state observed in the presence of chemical denaturants such as urea or guanidine hydrochloride. Recent studies have shown that TFE-state of a protein, e.g. as obtained by dissolving the protein (lysozyme, ctlactalbumin) in 50% (by vol.) aqueous TFE, is a stable, partially folded state with a high content of r conformation, but lacking the specific tertiary interactions of the native protein (Buck et al., 1993, 1995, 1996; Alexandrescu et al., 1994). This TFEstate appears to be a non-compact, expanded conformational state characterized by an ensemble of fluctuating helices. However, the TFE-state does not result from a gross structural reorganization of the protein to another unrelated structure, since the large majority of amides protected from exchange in the TFE-state are also protected in the native state of the protein, thus implying a similar location of helical segments along the polypeptide chain in both native and TFE-state (Buck et al., 1993, 1995, 1996; Alexandrescu et al., 1994). In recent years we have been interested at demonstrating that proteolyfic enzymes can be used as reliable probes of protein structure and dynamics (Fontana et al., 1986, 1993, 1997a,b). The outcome of these studies allowed us to propose that the key feature of the sites of initial proteolysis of a globular protein substrate resides in the enhanced flexibility (local unfolding) and, in particular, that helical chain segments are not prone to proteolysis (Polverino de Laureto et al., 1995a; Fontana et al., 1997a,b). On this basis, it was expected that the highly helical conformation of a protein substrate dissolved in aqueous TFE would hampers extensive proteolysis. We have conducted
383
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Figure 1. Far-UV (A) and near-UV (B) circular dichroism (CD) spectra of hen egg-white lysozyme (top) and thermolysin (bottom). The spectra of lysozyme were recorded at room temperature in 20 mM sodium phosphate buffer, pH 7,0, containing 50 mM NaCI and various amounts of TFE. The numbers near the curves indicate the % of TFE. In the case of thermolysin, the CD spectra were recorded in 20 mM Tris-HC1 buffer, pH 7.2, containing 5 mM CaCI2 in the absence ( ~ ) or presence of 50% TFE(-.-). Protein concentrations were about 0.1 and 0.8 mg/ml in the far- and near-UV regions, respectively. proteolysis of several model proteins in their TFE-state utilizing thennolysin (Fontana et al., 1995; Polverino de Laureto et al., 1995b, c; 1997). This thermophilic metalloendopeptidase (Matthews, 1988) appeared to be a most suitable proteolytic probe, because of its noteworthy stability under relatively harsh solvent conditions, including organic solvents (Welinder, 1988), and broad substrate specificity (Morihara & Tzusuki, 1970; Heinrikson, 1977; Keil, 1982). Thus, it was anticipated that thennolysin would cleave proteins in their TFE-state at sites characterized by the flexibility required for an
384
efficient proteolysis and not by the specificity of the protease. The striking observation emerged from these studies is that proteolysis by thermolysin in 50% TFE can be very selective, indicating that this novel procedure can be useful for preparing nicked proteins and/or large protein fragments. 2. PROTEINS IN 50% TFE The conformational properties of four model proteins (hen egg-white lysozyme, bovine pancreatic ribonuclease A, bovine ct-lactalbumin and horse cytochrome c) dissolved in aqueous TFE were examined by far-UV circular dichroism (CD) spectroscopy. As an example, Fig. 1 shows the CD spectra of lysozyme in aqueous buffer, pH 7.0, in the presence of increasing concentrations of TFE. The features of the far- and near-UV CD spectra clearly show that the action of alcohol is to enhance the helical secondary structure of the protein, as deduced by the shape of the CD spectra and ellipticity values at 208 and 280 nm, these last being diagnostic of helical polypeptides (Greenfield & Fasman, 1969; Johnson, 1990). It has been estimated that the four models proteins when dissolved in 50% TFE acquire a conformational state characterized by 45-60% helicity, compared with the 20-40% helical content in aqueous buffer (Fontana et al., 1995; Polverino de Laureto et al., 1995b,c; 1997; Galat, 1985; see also Shiraki et al., 1995). Conversely, the tertiary structure of lysozyme, as well as of the other model proteins, dissolved in 50% aqueous TFE is largely eliminated, as given by the strong reduction of the CD signal in the 250-300 nm region of the native protein (Stricldand, 1974) (see Fig. 1). Thus, from the CD data, it can be inferred that the TFEstate of proteins is characterized by a high helical secondary structure, but lacking the specific tertiary interactions of the native protein. Fig. 1 shows the CD spectra of thermolysin dissolved in 50% TFE at neutral pH. Clearly, thermolysin under these solvent conditions appears to maintain the integrity of its secondary and tertiary structure, since far- and near-UV CD spectra in buffer only and in 50% TFE are essentially identical. The unusual TFE-stability of thermolysin is in line with the fact that this exceptionally stable thermophilic enzyme can be crystallized from a 70% aqueous dimethylsulfoxide solution (Colman et al., 1974). 3. PROTEOLYSIS OF PROTEINS IN THEIR TFE-STATE Four model proteins have been reacted with thermolysin in 50% TFE at neutral pH and 20-52~ for several hours. Calcium (5-10 raM) was added to the reaction mixture, since this ion stabilizes thermolysin (Roche & Voordouw, 1978; Fontana et al., 1977). While all four model proteins (see above) were relatively resistant to proteolysis by thermolysin if incubated for a short time in buffer only and without TFE, proteolysis of these proteins in their TFE-state occurred slowly and very selective, as evidenced from the analysis of aliquots of the proteolytic mixture by sodium dodecyl sulphate (SDS) polyaerylamide gel electrophoresis (PAGE) and reverse-phase high
385
Figure 2. SDS-PAGE analysis of the proteolysis of lysozyme by thermolysin. (A) Proteolysis was conducted at 40~ in 50 mM Tris-HC1 buffer, pH 7.0, containing 5 mM CaCI2 in the presence of 50% TFE. The thermolysin:lysozyme ratio was 1:20 (by mass). Aliquots were taken from the reaction mixture alter 0, l, 2, 3, 4, 6 and 24 hours (lanes 1-7) and analyzed by SDSPAGE under reducing conditions (Schagger & vonJagow, 1957). (]3) SDS-PAGE analysis of purified nicked (lane l) and intact (lane 2) lysozyme. The nicked protein species is constituted by fragments 1-97 and 98-129 covalently linked by disulfide bonds. B
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386
Table 1. Limited proteolysis of proteins by thermolysin in 50% aqueous trifluoroethanol,a Protein
E:Sb (by mass)
Temperature (~
Peptide bond hydrolyzedc
Amino acid residuesa'''
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Slow
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25
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8
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52
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aThe protein was dissolved (~ 1 mg/ml) in 50 mM Tris-HCl buffer, pH 7.0, containing 5 mM CaC12 in the presence of 50% TFE. In the ease of cytochrome e, the buffer was 20 mM Tris, pH 7.8, containing 10 mM CaCI2. An aliquot of thermolysin, dissolved in the same buffer, was added to the protein solution and then the reaction mixture was kept at the indicated temperature for 0.5-24 hours. The time course of the proteolysis was determined by SDS-PAGE and reverse-phase HPLC. bThermolysin to protein ratio. CThe sites of proteolytic cleavage along the polypeptide chain of the protein were determined by N-terminal sequencing of the various fragments produced and comparing these data with the known amino acid sequence of the protein. The term fast and slow refers to the initial (after 1-3 hours) and subsequent (after 6-24 hours) peptide bond fission, respectively, dResidues per molecule. performance liquid chromatography (HPLC). As an example, the SDS-PAGE data obtained with lysozyme are shown in Fig. 2. It is seen that at 40~ lysozyme is slowly digested (1-6 hours of reaction) by thermolysin to two protein fragments only, while on prolonged reaction (24 hours) other fragments do appear in the Coomassie-stained gel. The thermolytic mixture of lysozyme (5 hours of reaction at 52~ has been analyzed also by reverse-phase HPLC, allowing us to isolate to homogeneity the major protein species (nicked protein) (Fig. 3A). A sample of nicked lysozyme, after reduction with excess thiol and S-carboxamidomethylation, eluted from the reverse-phase HPLC column in two chromatographic peaks (Fig. 3B). The protein material of these two peaks was fia~er analyzed in terms of N-terminal sequence and amino acid composition. These data, when compared with the known amino acid sequence of hen lysozyme, allowed us to establish that nicked lysozyme resulted from specific cleavage of the Lys97-Ile98 peptide bond and thus constituted by fragments 1-97 and 98-129 covalently linked by the four disulfide bridges of the protein. Moreover, it was found that upon prolonged proteolysis of lysozyme at high temperature, such as 24 hours at 40~ or 6 hours at 60~ additional but few cleavages of the protein do occur. In the case of lysozyme, these minor cleavages occurred at the Ser24-Leu25 and Asn37-Phe38 peptide bonds (Polverino de Laureto et al., 1995c).
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Figure 4. Sites of limited proteolysis along the amino acid sequences of horse cytochrome c, bovine pancreatic ribonuclease A, bovine a-lactalbumin and hen egg-white lysozyme. Major and minor arrows indicate the sites of fast and slow peptide bond fission, respectively (see also Table 1). The helical chain segments of the four proteins in their native state in aqueous buffer (without TFE), as given by crystallographic analysis of these proteins, are boxed. The selective proteolysis of lysozyme in its TFE-state by thermolysin (Polverino de Laureto et al., 1995c) parallels those observed with bovine a-lactalbumin (Polverino de Laureto et al., 1995b), horse cytochrome c (Fontana et al., 1995) and ribonuclease A (Polverino de Laureto et al., 1997). The results of the proteolysis experiments on the four model proteins are summarized in Table 1 and Fig. 4. Of interest, it was possible to isolate to homogeneity a nicked protein constituted by two fragments covalently linked by disulfide bridges in the case of lysozyme, r and ribonuclease A. The conformational, stability and functional properties of these nicked proteins were analyzed (Polverino de Laureto et al., 1995b,c; 1997). In the case of horse cytochrome c, the major thermolytic cleavage occurs at peptide bond Gly56-Ile57 (Fontana et al., 1995). The N-terminal fragment 1-56 maintains covalenfly bound the heine group of cytochrome c via a thioether linkage at the level of Cysl4 and Cysl7 and forms in
388
X-Leu = 100
lie
myr
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Ai~
THERMOLYSIN
---- Ser ---Thr
Figure 5. Relative rates of peptidr bond hydrolysis by thermolysin in aqueous buffer at neutral pH. The rates are graphically expressed as percent of that of the hydrolysis of the peptidr bond most preferentially cleaved by thermolysin. Of note, this metallo-cndopeptidasr cleaves at the amino side of bulky and hydrophobic amino acid residues (e.g., the peptide bond Xaa-Leu is cleaved). The data are taken from Keil (1982). aqueous buffer a non-covalent complex with the C-terminal fragment 57-104 (A. Fontana et al., unpublished). 4. DISCUSSION The results of this study show that a protein dissolved in aqueous TFE can be cleaved by thcrmolysin at very few pcptidc bonds, leading to quite large protein fragments and/or nicked protein species rather resistant to further protcolysis. The ability of proteins in their helical TFE-statc to resist extensive protcolytic degradation appears to be remarkable, if one considers the broad substratc specificity of thermolysin in aqueous buffer at neutral pH (Morihara & Tzusuld, 1970; Hcinrikson, 1977; Kcil, 1982). As shown in Fig. 5, thcrmolysin in water cleaves mostly at the amino side of bulky and hydrophobic amino acids, but si~,nificant hydrolysis occurs at other residues as well. On the other hand, the data given in Table 1 and Fig. 4 indicate that in aqueous TFE both major and minor cleavages by thennolysin of the four model proteins herewith investigated occur at pcptidc bonds involving only Ilc, Lcu and Phc residues. Nevertheless, even if the model proteins contain many Lcu, Ilc and Phc residues (see Fig. 4), thcrmolysin cleaves these proteins at very few sites only. With each model protein examined there is a pcptidc bond being cleaved first by thermolysin, followed by much slower cleavages at few additional peptidc bonds. The selective cleavage of proteins by thcrmolysin in aqueous TFE must be dictated both by the conformational state of the protein substratc and by some specific features of the biocatalyst dissolved in aqueous TFE. First of all, the protein substratc acquires a high hehcal state in TFE, since this alcohol appears to favour the formation of intramolccular hydrogen bonds of the ct-hchcal conformations rather than hydrogen bonds with the solvent (Thomas & Dill, 1993; Jasanoff & Fcrsht, 1994; CammcrsGoodwin ct al., 1996; Luo & Baldwin, 1997; Myers ct al., 1998). Dobson and co-
389 workers (Buck et al., 1993, 1995, 1996; Alexandreseu et al., 1994; Affleek et al., 1994) demonstrated that at least the helical segments of the native protein are maintained in the TFE-state, even if hydrogen exchange measurements indicate that these helices are highly dynamic entities (fluctuating). Moreover, TFE also induces the formation of some extra helices which do not correspond to native-like helices (Buck et al., 1993, 1995, 1996; Alexandreseu et al., 1994). Thus, the high helical state of the protein substrate in aqueous TFE is not expected to be easily hydrolyzed by the protease, since a folded protein molecule is much more resistant to proteolysis than a random-coil polypeptide chain (Mihalyi, 1978). That the helical TFE-state of proteins is hampering extensive proteolysis is also in line with our proposal that proteolysis does not occur at the level of helical chain segments and that only flexible loops are the sites of fast proteolysis (Fontana et al., 1986, 1993, 1997a, b; Polverino de Laureto et al., 1995a). The selective proteolysis of proteins in aqueous TFE by thermolysin occurs rather slowly, if one considers that proteolysis requires several hours of incubation at moderately high temperatures (e.g., 25-52~ see Table 1). This is due not only to the structured (helical) protein substrate hampering binding and adaptation at the active site of the protease, but also to the fact that the organic solvent causes significant enzyme inhibition (see Klibanov, 1997, for a recent discussion). It is tempting to speculate that, in the presence of the organic cosolvent, thermolysin can acquire an enhanced (overall and local) protein rigidity (Affleck et al., 1992a, b; Hartsough & Merz, 1992) causing a reduction of its catalytic potency, since some chain mobility is required for catalysis (Welch, 1986; Fersht, 1985; Kraut, 1988). The fact that thermolysin is much less active in the presence of TFE is also explained by the fact that proteases, including thermolysin (Wayne & Fruton, 1983), in the presence of organic cosolvents catalyze the reverse reaction, i.e. the synthesis instead of the hydrolysis of the peptide bond (Chaiken, 1981; Kullman, 1987). The proteolysis of proteins in their TFE-state can be interpreted also on the basis of some features of the structure and dynamics of the native protein in aqueous solution, implying that the helical TFE-state appears to be related to the native state. First of all, as shown in Fig. 4, the sites of initial cleavages within the polypeptide chains of the four model proteins occur outside the helical chain segments of the native proteins. Accepting our view that helices are quite rigid rods not prone to proteolysis (Fontana et al., 1986, 1993, 1997a, b; Polverino de Laureto et al., 1995a), proteins in their TFE-state appear to maintain at least the helical chain segments of their native state. Moreover, the sites of the proteolytic cleavage along the protein chain (Fig. 4) should be the most flexible ones, since we have previously demonstrated that there is a correlation between site: of limited proteolysis and sites of higher segmental mobility (Fontana et al., 1986). For example, the chain region encompassing the Asn34-Leu35 peptide bond which is cleaved in ribonuclease A in its TFE-state (see Fig. 4) is the most flexible one also in the native protein in aqueous buffer, as given by hydrogen exchange measurements (Kiefhaber & Baldwin, 1995) and by the fact that this region is poorly defined in the Xray density map (Wlodawer et al., 1982). Of interest, the same Asn34-Leu35 peptide bond is the site of initial hydrolysis when ribonuclease is reacted with thermolysin on
390 mild heating in aqueous buffer (Arnold et al., 1996). All these data therefore indicate that the chain region encompassing the Asn34-Leu35 peptide bond in ribonuclease A displays higher flexibility than the rest of the protein chain both in the native state and in TFE-state, implying some similarity of structure and dynamics between the two states. In summary, proteolysis of proteins in aqueous TFE by thermolysin is a novel and useful procedure for the selective enzymatic fragmentation of proteins. The technique is simple to use, modest in demands for protein sample and experimental effort and will fred useful applications in protein structure research for the preparation of rather large protein fragments for sequencing purposes by the Edman technique and for biophysical and functional studies. A..eknowledgements. This study was supported in part by the Italian National Council of Research (CNR), special project on Biotechnology, and by the Ministery of the University and Scientific Research (MURST). 5. REFERENCES Affleck, A.T., Ng, Y.-L. & Dobson, C.M. (1994)J. Mol. Biol. 235, 587-599. Affleek, R., Hayness, C.A. & Clark, D.S. (1992a) Proc. Natl. Acad. ScL USA 89, 51675170. Affleck, R., Xu, Z.F., Suzawa, V., Focht, K., Clark, D.S. & Dordick, J.S. (1992b) Proc. Natl. Acad. Sci. USA 89, 1100-1104. Alexandrescu, A.T., Ng, Y.-L. & Dobson, C.M. (1994)3'. Mol. Biol. 235, 587-599. Arnold, V., Rticknagel, K.P. Schirhom, A. & Ulbrick-Hofman, R. (1996) Fur. jr. Biochem. 237, 862-869. Bolin, K.A., Pitkeathly, M., Miranker, A., Smith, L.J. & Dobson, C.M. (1996)Jr. Mol. Biol. 261,443-453. Buck, M., Radford, S.E. & Dobson, C.M. (1993) Biochemistry 32, 669-678. Buck, M., Schwalbe, H. & Dobson, C.M. (1995) Biochemistry 34, 13219-13232. Buck, M., Schwalbe, H. & Dobson, C.M. (1996)3'. Mol. Biol. 257, 669-683. Cammers-Goodwin, A., Allen, J.J., Oslick, S.L., McLure, K.F., Lee, J.H. & Kemp, D.S. (1996) or. Amer. Chem. Soc. 118, 3082-3090. Carrea, G., Ottolina, G. & Riva, S. (1995) Trends Biotechnol. 13, 63-70. Chaiken, I.M. (1981) CRC Crit. Rev. Biochem. 11,255-301. Colman, P.M, Jansonius, J.N. & Matthews, B.M. (1974)J. Mol. Biol. 70, 701-724. De Filippis, V. & Fontana, A. (1990) Int. J. Peptide Protein Res. 35, 219-227. Dordick, J.S. (1989)Enzyme Microb. Technol. 11, 194-211. Fan, P., Bracken, C. & Baum, J. (1993) Biochemistry 32, 1573-1582. Fersht, A. (1985) Enzyme Structure and Mechanism, 2nd ed., pp. 293-369, Freeman, San Francisco. Fontana, A., Fassina, G., Vita, C., Dalzoppo, D., Zamai, M. & Zambonin, M. (1986) Biochemistry 25, 1847-1851.
391 Fontana, A., Polverino de Laureto, P. & De Filippis, V. (1993) in Stability and Stabilization of Enzymes (van den Tweel, W.J.J., Harder, A. & Buitelaar, R.M., eds.), pp. 101-110, Elsevier, Amsterdam. Fontana, A., Zarnbonin, M., Polverino de Laureto, P., De Filippis, V., Clementi, A. & Scaramella, E. (1997a)J. Mol. Biol. 266, 223-230. Fontaaa, A., Polverino de Laureto, P., De Filippis, V., Scaramella, E. & Zambonin, M. (1997b) Folding & Design 2, R17-R26. Fontana, A., Vita, C., Bocc/~, E. & Veronese, F.M. (1977) Biochem. J. 165, 539-545. Fontana, A., Zambonin, M., De Filippis, V., Bosco, M. & Polverino de Laureto, P. (1995) FEBS Lett. 362, 266-270. Galat, A. (1985) Bochim. Biophys. Acta 827, 221-227. Greenfield, N.J. & Fasman, G.D. (1969)Biochemistry 8, 4108-4116. Gupta, M.N. (1992)Eur. ,1. Biochem. 203, 25-32. Hamada, D., Kuroda, Y., Tanaka, T. & Goto, Y. (1995)Jr. Mol. Biol. 254, 737-746. Hartsough, D.S. & Merz, K.M. (1992),/. Amer. Chem. Soc. 114, 10113-10116. Heinrikson, R.L. (1977) Methods Enzymol. 3, 531-568. Jasanoff, A. & Fersht, A.R. (1994) Biochemistry 33, 2129-2135. Jayaraman, G., Kumar, T.K.S., Anmkamaar, A.I. & Yu, C. (1996) Biochem. Biophys. Res. Commun. 222, 33-37. Jencks, W.P. (1969) in Catalysis in Chemistry and Enzymology, pp. 321-436, McGraw Hill, New York. Johnson, W.C. (1990) Proteins Struct. Funct. Genet. 7, 205-214. Keil, B. (1982) in Methods in Protein Sequence Analysis (Elzinga, M., ed.) pp. 291-304, Humana Press, Clifton, New York. Kieflaaber, T. & Baldwin, R.L. (1995) Proc. Natl. Acad. ScL USA 92, 2657-2661. Klibanov, A.M. (1989) Trends Biochem. Sci. 14, 141-144. Klibanov, A.M. (1997) Trends Biotechnol. 15, 97-101. Kraut, J. (1988) Science 242, 533-540. Kullman, W. (1987) Enzymatic Peptide Synthesis, CRC Press, Boca Raton, Florida. Lehnnan, S.R., Tuls, J.L. & Lund, M. (1990) Biochemistry 29, 5590-5596. Lign6, T., Pauthe, E., Monti, J.P., Gacel, G. & Larreta-Garde, V. (1997) Biochim. Biophys. Acta 1337, 143-148. Luo, P. & Baldwin, R.L. (1997) Biochemistry 36, 8413-8421. Matthews, B.W. (1988)Acc. Chem. Res. 21, 333-340. Mattiasson, B. & Adlercreutz, P. (1991) Trends Biotechnol. 9, 394-398. Mih~yi, E. (1978) Application of Proteolytic Enzymes to Protein Structure Studies, CRC Press, Boca Raton, Florida. Morihara, K. & Tzusuki, H. (1970) Eur. J. Biochem. 15, 374-380. Myers, J.K., Pace, C.N. & Scholtz, J.M. (1998) Protein ScL 7, 383-388. Nelson, J.W. & Kallenbach, N.R. (1986) Proteins Struct. Funct. Genet. 1, 211-217. Polverino de Laureto, P., Toma, S., Tonon, G. & Fontana, A. (1995a) Int. J. Pepade Protein Res. 45, 200-208.
392 Polverino de Laureto, P., De Filippis, V., Di Bello, M., Zambonin, M. & Fontana, A. (1995b) Biochemistry 34, 12596-12604. Polverino de Laureto, P., De Filippis, V., ScarameUa, E., Zambonin, M. & Fontana, A. (1995c) Fur. d. Biochem. 230, 779-787. Polverino de Laureto, P., Scaramella, E., De Filippis, V., Bmix, M., Rico, M. & Fontana, A. (1997) Protein Sci. 6, 860-872. Pourplanche, C., Lambert, C., Beryot, M., Minx, J., Chopard, C., Alix, A.J.P. & Larreta-Gardr V. (1994)s Biol. Chem. 269, 31585-31591. Roche, R.S. & Voordouw, G. (1978) CRC Crit. Rev. Biochem. 5, 1-23. Schiigger, H. &von Jagow, G. (1987) Anal. Biochem. 166, 368-379. Segawa, S.I., Fukuno, T., Fujiwara, K. & Noda, Y. (1991) Biopolymers 31, 497-509. Shiraki, K., Nishikawa, K. & Goto, Y. (1995)d. Mol. Biol. 245, 180-194. Smith, L.J., Alexandrescu, A.T., Pitkeathly, M. & Dobson, C.M. (1994) Structure 2, 703-712. SOnnichsen, F.D., Van Eyk, J.E., Hodges, R.S. & Sykes, B.D. (1992) Biochemistry 31, 8790-8798. Storrs, R.W., Truckses, D. & Wemmer, D.E. (1992) Biopolymers 32, 1695-1792. Strickland, E.H. (1974)CRC Crit. Rev. Biochem. 2, 113-174. Tamburro, A.M., Scatmrin, A., Rocchi, R., Marchiori, F., Borin, G. & Scoffone, E. (1968) FEBS Lett. l, 298-300. Thomas, P.D. & Dill, K.A. (1993) Protein Sci. 2, 2050-2065. Wayne, S.I. & Fruton, J.S. (1983) Proc. Natl. Acad. Sci. USA 80, 3241-3244. Welch, G.R. (1986) The Fluctuating Enzyme, J. Wiley, New York. Welinder, K.G. (1988) Anal. Biochem. 174, 54-64. Wlodawer, A., Bott, R. & Sj61in, L. (1982)d. Biol. Chem. 257, 1325-1332. Wong, C.-H. (1989) Science 244, 1145-1152.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
393
Enzyme inactivation by inert gas bubbling Myl6ne Caussette, Alain Gaunand, Henri Planche and Brigitte Lindet Ecole Nationale Sup&ieure des Mines de Paris, CEREP, Biotechnology Laboratory, 60, Boulevard Saint-Michel, 75006 Paris, France
Nitrogen bubbling strongly enhances the inactivation of enzymes such as lysozyme, lipase or pectinmethylesterase.The nitrogen bubbling efficiency depends on the enzyme. The inactivation kinetics are first order except for lipase. A rapid initial phase is followed by a decrease of the inactivation rate which has been explained by a competition between active and inactive lipase species at the gas-liquid interface. The inactivation kinetics of lysozyme and lipase increase linearly with the specific interfacial area. For pectinmethylesterase the inactivation kinetics do not depend on the specific interfacial area. For the three enzymes, physico-chemical parameters like pH and ionic strength act on the inactivation efficiency by probably modifying the adsorption mechanisms. This study points out the importance of hydrodynamics in bioreactors (loss of enzyme activity during their production by fermentation or loss of enzyme efficiency during biocatalysis reactions). This work also highlights the potentialities of an inactivation process using gas-liquid interfaces.
1. INTRODUCTION Experimental observations in our laboratory have shown that gas bubbling induced enzyme inactivation (1). The interest of this observation relies on the two following factors. On the one hand, to reach high yields ofbioconversion and high productivities, it is essential to control the parameters acting on enzyme stability. On the other hand, in industrial processes, enzyme inactivation is generally required to stop the enzymatic reaction and to prevent product deterioration which can affect the final quality of the product. Very few works correlate enzyme inactivation and the presence of interfaces. Some studies have focused on enzyme adsorption at solid-liquid interfaces (2-5). At liquid-liquid interfaces, enzyme inactivation has been mainly reported by the Pr. Halling's team (6-9). Some works have reported the protein adsorption at static air-water interface (10-16). Enzyme inactivation at this interface has only been reported by few authors (17,18). This article focuses on enzyme inactivation at dynamic gas-liquid interfaces. Inactivation studies have been performed by measuring residual enzymatic activity after nitrogen bubbling in an enzyme aqueous solution. Three enzymes have been studied : Lysozyme, Lipase and Pectinmethylesterase.
394 2. MATERIALS AND METHODS
2.1. Enzymes Chicken egg-white lysozyme was supplied by Sigma Chemical Co (St Louis Mo), crystallised three-times, dialysed and lyophilised. Lipase from Aspergillus oryzae was supplied by NovoNordisk,(Lipolase) and Pectinmethylesterase, (PME) from Aspergillus niger was provided by Gist-Brocades. PME showed a single band on agarose gel electrophoresis. 2.2. Substrates and chemicals Micrococcus lysodeikticus (Sigma) was used as the lysozyme substrate for activity measurements. Tributyrin, lipase substrate, came from ICN Biomedicals and gum arabic from Prolabo. Pectin (SBI, France) with a methylation degree > 70 % was used as the PME substrate. All the other chemicals used in this work were of the highest grade commercially available. Nitrogen was from Airgaz (France) with a certified purity of 99.995 %. 2.3. Bubble column The 0.015 m internal diameter and 0.4 m height column was thermostated with a double coated envelope.Pure nitrogen from a gas cylinder was bubbled in the column through a gas sparger located at its bottom equiped by three single nozzles 5x105 m in diameter.The nitrogen flow rate was controlled with a mass flow meter calibrated between 0 and 400 ml.min t with an error < 0.1%. The gas was humidified and preheated to working temperature in order to avoid water evaporation and temperature variation.The effect of gas sparging was tested at several flow rates ranging from 14 ml.min ~ to 150 ml.min ~. All the assays were done for a 60 ml enzyme solution. 2.4. Enzyme activity measurement Lysozyme activity was determined at 25C by measuring the decrease in absorbance at 450 nm of a M. lysodeikticus suspension 0.16 g.Lt, in 0.1 M potassium phosphate buffer pH 6.2. Lipase activity was determined by measuring at 30~ and pH 7, the release of fatty acids coming from the hydrolysis of a 0.05 % tribut~n emulsion. The PME activity was determined at 25~ and pH 4.5 by automatic titration according to the method of Rouse and Atkins (19). 2.5. Inactivation experiments Enzyme was solubilized in a preheated buffer solution, (phosphate buffer 0.01M for lysozyme and lipase, and citrate buffer 0.01M for PME). Enzymatic solution (60 mL) was stirred by the bubble rise in the column. No additional mechanical stirrer was used. Because of differences in the enzymes thermostability, the inactivation kinetics have been performed at different temperatures. At different times, samples were withdrawn from the solution. Each sample was immediately cooled in a O~ water bath in order to stop the inactivation process. The enzyme activity was then measured according to the previously reported methods. Results are expressed as a percentage of residual activity versus time. The residual activity is defined as
395 the ratio of the activity at time t to the initial activity. The half-life is the time when the enzyme lost 50% of its activity.
3. RESULTS 3.1. Effect of nitrogen bubbling on enzyme inactivation Inactivation is strongly enhanced by gas bubbling for lysozyme, lipase and PME as shown on figures 1 and 2.
Fimtre 1: Influence of nitrogen bubbling on the inactivation kinetics of lysozyme and lipase Lysozyme:4~ :Q=0ml.min ~;m:Q=100ml.mn -I Lipase: A:Q=0ml.min~; :~:Q=100ml.mn ~ Experimental conditions for lysozyme experiments :Phosphate buffer 0.01M pH 6.2; Lysozyme concentration : 0.05 g.L~; Temperature: 70~ Experimental conditions for lipase experiments : Phosphate buffer 0.01M pH 6.2; Lipase concentration: 0.02 g.L~; Temperature: 80~
Figure 2" Influence of nitrogen bubbling on PME inactivation kinetics PME:m :Q=0 ml.mn 1 PME: .~.:Q=100ml.mn-~ Experimental conditions: Citrate buffer 0.01M pH 4.5; PME concentration : 0.008 g.L~;Temperature 50~
As an example, for a 100 mL.min"1flow rate at 70~ the half life of lysozyme is 15 minutes while in the same conditions without gas bubbling no activity loss is observed after 8 hours. For the same flow rate at 80~ the half life of lipase decreases from 3 hours without nitrogen bubbling to less than 5 minutes with bubbling. However gas bubbling is less efficient on PME for which the half life decreases from 8.5 hours to 3.5 hours at 50~ In the first stages of the inactivation process, the inactivation kinetics are first order for the three enzymes. For lipase, a change of the inactivation kinetic order appears during the inactivation process. No
396 reactivation was observed when samples were incubated for a further 24 hours without bubbling. Gas promoted inactivation is an irreversible process.
3.2. Effect of lipase inactivated species. Further experiments have been performed to explain lipase behaviour. The hypothesis of a competition at gas-liquid interface between active and inactive species has been made. Inactivation of lipase by nitrogen bubbling was carried out with and without inactivated lipase molecules at initial time, (these molecules were inactivated before by heat treatment). The resulting inactivation kinetics are plotted on figure 3. In the presence of inactivated lipase molecules at initial time, a decrease of the inactivation kinetics is observed. A control experiment without bubbling shows that the presence of inactivated lipase species do not affect inactivation. Competition effect at gas-liquid interfaces between active and inactive species might explain lipase behaviour.
90-
g 80 ,~ 70 g9 60
4,
,e,
,e,
I
I
i
. . .
o m tl = I/) | o, ,,,.,
50 40 30 20 10 0
9 =
0
P'
10
I
I
20 30 T i m e (rain)
I
'1'
40
50
a I
60
Figure 3: Influence of inactivated species on lipase inactivation kinetics O:Q =0 ml.min-1, I = 0 g.H ;l:Q=100ml.rnn-1, I=O g.N ;A:Q= 100 ml.min -1, I=0,02 g.N Experimental conditions: Phosphate buffer 0.01M p H 6.2; Initial concentration of active lipase 90.02 g.L-~; Initial concentration of thermo-inactivated lipase" 0 and 0.02 g.L-1; Temperature" 80~
-v -v 100 "~ -v" = A 90 #80 9 9 ,~, 70 9 7_> 60 X ,,i.,a o 50 & 40 Ill X ,It& = 30 & 9 -~ 20 w • 9 9 10 iv, Xxx a, I 0 0 80 20 40 60 T i m e (rain)
9
I
100
120
Figure 4: pH influence on lysozyme inactivation kinetics 9 :pH = 4 ;m:pH = 6,2 ;A:pH = 7 ; :....:pH ---8. Experimental conditions" Phosphate buffer 0.01M; Lysozyme concentration" 0.05 g.L-~; Temperature- 70~ Nitrogen flow rate 950 mL.min-1
3.3. Effect of physico-chemicai parameters on enzyme inactivation kinetics. Several authors have demonstrated that physico-chemical parameters such as pH and ionic strentgh act on protein adsorption processes (5,20). Enhancing protein adsorption process might result in an increase of inactivation kinetics at gas-liquid interface. Figure 4 gives the pH effect on lysozyme inactivation at a 50 mL. min ~ nitrogen flow rate. Gas bubbling efficiency strongly depends on pH. A pH close to the enzyme isoelectric point enhances the
397 inactivation. No activity loss is observed for experiments carded out without bubbling in the same conditions of pH, temperature and duration. The same kind of effect has been observed for the other enzymes (data not shown). Table 1 shows the influence of NaC1 concentration on PME half life with and without bubbling. Without bubbling, NaC1 exerts a protective effect on PME. The PME half life increases with NaC1 concentration. With bubbling, PME half life decreases very strongly. At 2M NaCI, the PME half life decreases from 91 min to 9.5 min when bubbling at 100 mL. min-~" Table 1 Effect of NaC1 concentration on PME inactivation NaC1 (M) 0
1
2
3
Half life (min), Q = 0 mL.min~
62
101
91
95
Half life (min), Q= 100 mL.min ~
48
53
9,5
5,6
Experimental conditions: PME concentration 0,008 g.L~; Citrate buffer 0,01M; pH 4,5; Nitrogen flow rate : 0 and 100 mL.min~; Temperature : 54.5~
4. DISCUSSION Inactivation is strongly enhanced by gas bubbling for the three studied enzymes, lysozyme, lipase and pectinmethylesterase. Nitrogen bubbling in an enzyme solution generates irreversibly inactivated enzyme molecules. For the studied enzyme concentrations, the inactivation kinetics of lysozyme and PME in the presence of nitrogen bubbles have been found of first order. For lipase, the change of the reaction order throughout the inactivation process can be explained by the formation of inactivated lipase species. Inactive lipase species induce a competition effect at gas-liquid interfaces between active and inactive species which consequently slows down lipase inactivation kinetics. Efficiency of gas bubbling on enzyme inactivation strongly depends on pH and ionic strentgh. The same tendency was observed for the three enzymes : the closer the pH is to the enzyme isoelectric point (pI) the more efficient the inactivation process is. (pI of lysozyme, lipase and PME are respectively 11, 4.3 and 4.4). The pH and ionic strength act on the enzyme adsorption process and enhance the inactivation kinetics. Our results seem to demonstrate that inactivation of lysozyme, lipase and PME promoted by gas bubbling is mainly governed by an interfacial process.
398 REFERENCES
1. 2. 3. 4. 5. 6. 7.
Lindet B., Caussette M., Delepine S. and Planche H. (1995). Patent FR 9515239. Norde W., Adv. Coll. Int. Sci., 25 (1986) 267-340. Schmidt C.F., Zimmermann R.M., Gaub H.E., Biophys. J., 54 (1990) 577-588. Ball A., Jones R.A.L., Langrnuir, 11 (1995) 3542-3548. Buijs J., Norde W., Langmuir, 12 (1996) 1605-1613. Cassells J.M., HaUingG P.J., Enzyme Microb. Technol., 12 (1990) 755-759. Blanco R.M., Hailing P.J., Bastida A., CuestaA C., Guisan J.M., Biotechnol. Bioeng., 39 (1992) 75-84. 8. Ghatorae A.S., Guerra M.J., Bell G. Hailing P.J., Biotechnol. Bioeng., 44 (1994) 13551361. 9. Ghatorae A.S., Bell G. Hailing P.J., Bioteehnol. Bioeng., 43 (1994) 331-336. 10. Adams D.J., Evans M.T.A., Mitchell J.R., Phillips M.C., Rees P.M., J. Polymer Sci., Part C, 34, (1971) 167-179. 11. Graham D.E., Phillips M.C., J. Coll. Int. Sci., 70, 3 (1979) 403-439. 12. Graham D.E., Phillips M.C., J. Coll. Int. Sci., 76, 1 (1980) 240-249. 13. Ter-Minassian-Saraga L., J. Coll. Int. Sci., 80, 2 (1981) 393-401. 14. Hunter J.R., Carbonell R.G. Kilpatrick P.K., J. Coll. Int. Sci., 143, 1 (1991) 37-53. 15. Mac Ritchie F., Anal. Chim. Acta, 249 (1991) 241-245. 16. Tronin A., Dubrovsky T., Dubroskaya S., Radicchi G. and Nicolini C., Langrnuir 12 (1996) 3272-3275. 17. Inbar I. and Miller, I., Biochim. Biophys. Aeta, 452 (1976) 417-430. 18. Xu S. and Damodaran S., J. Coll. Int.Sci. 159 (1993) 124-133. 19. Rouse, A. H. and Atkins, C., Food Teehnol 6: (1952) 291-294. 20. Wannerberger K., Amerbrant T., J. Coll. Int. Sci., 117 (1996) 316-324.
Stability and Stabilization of Biocatalysts A. BaUesteros, F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Effects of water-miscible solvents on the stability and specificity of cyclodextringlucosyltransferases Anne D. Blackwood and Christopher Bucke School of Biosciences, University of Westminster, 115 New Cavendish Street, London W lM 8JS, UK 1. INTRODUCTION. Cyclodextringlucosyltransferase (EC 3.2.1.19 [CGTase]) catalyses several reactions with starch as substrate, including the synthesis of three cyclodextrins(CD) (t~,13 and ?) with 6,7 and 8 glucose units joined by a-1,4 linkages. It also catalyses three transferase reactions, the hydrolysis of starch to linear dextrins, the disproportionation of two linear dextrins into linear oligosaccharides of different sizes and the coupling of CDs with linear dextrins. Several species and strains of bacteria produce CGTases all of which appear to be capable of catalysing all the reactions but which differ in their specificities. The CDs are the commercially important products of CGTase action: the separation and purification of the individual cyclodextrins is an expensive part of the production procedure. Consequently there is a need to find methods of improving the selectivity of CGTases to produce improved yields of individual CDs. Cyclodextrin yield and specificity can be improved by the inclusion in reaction mixtures of complexants which form insoluble selective inclusion complexes with CD molecules (1,2). There is also mention in the literature of increases in CD production achieved by the inclusion of low M.W. polar solvents such as ethanol (3) and acetone (4). The mechanism by which this effect occurs is less clear. To extend knowledge of this phenomenon, the effects of a range of organic co-solvents on the activities of CGTases from a Thermoanaerobacter sp. (5) and Bacillus circulans strain 251 (6) have been investigated. Both strains produce predominantly ~-CD in normal process conditions but the enzymes have been found to differ in their responses to the organic co-solvents used. 2. MATERIALS AND METHODS. 2.1. Enzymes and Chemicals. T-CGTase (from Thermoanaerobacter sp.) with a specific activity of 120 NU/g (based on its a-amylase activity measured by the Phadebas method) was a gift of Novo-Nordisk. B-CGTase (from Bacillus circulans strain 251) with a specific activity (for 13-CD production) of 270 U/mg was provided by Professor Lubber Dijkhuizen,
399
400 University of Groningen. All anhydrous organic solvents were stored over 4-A molecular sieves. Phadebas tablets (cross-linked insoluble blue-dyed starch) were from Pharmacia. Paselli SA2, a 2 dextrose equivalent maltodextrin preparation, was from Avebe, the Netherlands. 2.2. Enzyme Stability. T-CGTase was lyophilized from 0.1M Na phosphate buffer pH 6.0. Freeze-dried enzyme (lmg/ml) was suspended in 10 ml of an aqueous-organic solvent solution (aqueous phase was 0.1M Na-phosphate buffer pH 6.0) and incubated at for 24 h at 25~ Samples were then diluted 1000-fold and assayed for hydrolytic activity for 15 min at pH 6.2 and 60~ by measuring the absorbance at 620 nm of the blue dye released from Phadebas tablets. Enzyme activity was compared with a standard czamylase from Bacillus licheniformis. 2.3. Enzyme assays. Synthesis of I3-CD was followed using Paselli SA2 (5% w/v) as substrate by the formation of a colourless inclusion complex with phenolphthalein. One unit of activity produces 1~tmol of 0-CD per min. Solvent effects on CGTase activity were determined by incubating 0.2 U/ml of enzyme with 5%(w/v) Paselli SA2 in 10raM Na-citrate buffer, pH 6.0, at 50~ for 24 h. Samples were taken at regular time intervals, added to 100 B1 of 1M HC1 and boiled for 5 rain. Products were analysed by HPLC using a Dionex PAl anion-exchange column eluted with 150mM NaOH with Na acetate gradient and a flow rate of lml/min. Detection was by pulsed amperometric detector (PAD). The coupling activity of T-CGTase was measured in the presence of various additives (2.5 g/1 maltose and/or 10% v/v solvent) using either cx or ~3-CD as donor. T-CGTase (0.2U/ml) was added to the reaction in 10raM Na-citrate buffer, pH 6.0 and the residual CD after 4 h incubation at 50~ determined by HPLC. 3. RESULTS. The stability of the Thermoanaerobacter sp CGTase on exposttre to 12 polar organic solvents at a range of aqueous-organic solvent concentrations is summarised in Table 1. At low solvent concentrations the enzyme retained over 70% of its hydrolysis activity relative to enzyme incubated in the absence of organic solvent. At higher solvent concentrations T-CGTase was inactivated only in protein-dissolving solvents such as DMSO and in DMF, diethylacetamide (DMA) and pyridine. Table 1 provides also the partitioning coefficients, log P, for each solvent as measured in a standard octanol-water two-phase system Log P has been suggested as a reliable predictor ofbiocatalyst function in organic solvents (7): solvents with high log P values generally give the best retention of enzyme activity, while hydrophilic solvents with log P less than 2 are thought to remove the hydration "shell" of the enzyme, thus eliminating biocatalytic activity. DMSO and DMF both significantly denature TCGTase and have low log P values but other polar solvents of similar hydrophilicity do not denature the enzyme under these conditions. The enzyme maintained max~um activity, with even a slight increase in activity in all solvents at 20% (v/v) solvent concentration. This encouraged the investigation of solvent effects on enzyme specificity.
401 The effects of 8 polar solvents at 10% (v/v) concentration on the total conversion of Paselli SA2 to CDs by both T-CGTase and B-CGTase was examined. The results are summarised in Table 2. In all of the solvents tested there was an increase in the overall conversion of maltodextrin to cyclodextrin products compared with a solvent-free control. Maximum conversion, 66%, was achieved with the addition of 10% (v/v) ethanol. Addition of DMSO, DMA and t-butanol as co-solvents produced an TABLE 1. Log P values of solvents and relative residual hydrolysis activity ofTCGTase after 24 h incubation in solvent-aqueous buffer mixtures at 24~ % residual activity as a function of organic solvent concentration (v/v) Solvent
logP
20
40
50
70
100
Triethylene glycol dimethyl ether Dimethylsulfoxide Dioxane N,N-Dimethylformamide N,N-Dimethylacetamide Acetonitrile Ethanol Acetone Ethylene glycol diethyl ether Tetrahydrofuran Pyridine t-Butanol
- 1.86
107
99
99
93
77
- 1.3 - 1.1 - 1.0 -0.81 -0.33 -0.24 -0.23 0.28
108
87 80 86 70 87 100 97 98
81
75
104 83 98 92 105 118
103 87 93 83 91 90 95 100
3 22 82 89 94 108
7 80 72 74 61 64
0.49 0.71 0.78
102 82 100
85 78 91
77 71 96
73 27 90
66 57 70
104
0
13
increase in [3-CD formation with corresponding decreases in the ratios of both a and yCDs. DMF and acetone also produced increased synthesis of ~-CD with a decrease in ~/-CD production. Addition of acetonitrile, ethanol or tetrahydrofuran resulted in an increase in r formation at the expense of both [3 and y-CDs. The product selectivity of B-CGTase, determined under similar conditions is shown in Table 3. As with T-CGTase, each solvent produced an increase in the overall conversion of Paselli SA2 to CDs but the increases were less pronounced. Maximum conversion of maltodextrins to CDs (51%) was achieved with DMF as co-solvent. In contrast to TCGTase, each case addition of co-solvent resulted in an increase in the formation of [3CD at the expense of both ~ and ~/-CD. In other experiments (data not shown) it was demonstrated that with T-CGTase the inclusion ofjust 2% (v/v) solvent in the reaction mixture caused an increase in either r (acetonitrile) or [~-CD (DMA). Production of the favoured CD increased further with increases in co-solvent concentration up to 20% (v/v). The reason for this increase in CD production in the presence of small concentrations of polar organic solvents is unclear.
402 TABLE 2. Conversion of maltodextrins to cyclodextrins and product specificity ofTCGTase after 24 h incubation in 10% v/v organic solvent-buffer mixtures. Product ratio (%) Solvent
Substrate conversion to CDs (%)
r
13
7
None Dimethylsulfoxide Dimethylformamide Dimethylacetamide Acetonitrile Ethanol Acetone Tetrahvdrofuran
40 50 47 57 54 66 55 57
30 24 32 16 58 51 30 48
50 66 57 68 32 39 64 43
20 10 10 16 10 10 6 9
TABLE 3. Conversion ofmaltodextrins to cyclodextrins and product selectivity of BCGTase after 24 h incubation in 10% (v/v) organic solvent-aqueous buffer mixtures. Product ratio (%) Substrate conversion Solvent
to CDs (%)
~
13
None Dimethylsulfoxide Dimethylformamide Acetonitrile Tetrahydrofuran t-Butanol
40 45 51 45 42 44
15 11 9 7 10 9
65 77 77 76 76 82
20 12 14 16 14
9
No precipitation of product was observed in any of the organic co-solvent-aqueous buffer mixtures. It is plausa'ble that the solvents affect reactions which consume the CDs after synthesis or otherwise limit yields. In the conditions of industrial production of CD, yield is limited by product inhibition (8). In B-CGTase, which has five structural domains (6), [3-CD causes inhibition by interfering with the function of a maltose binding site located in the raw starch-binding E-domain of the enzyme as well as interfering with catalysis at the active site (9). The CD bound to the maltose binding site could interfere with the guidance of linear maltodextrin chains from this binding site to the active site located in the A-domain of the enzyme (6,10). Solvent interacting with the CD could alter the conformation of the CD and hinder or prevent its binding with the maltose binding site and/or the active site. It was found that addition of 0.2 mM I3-CD to T-CGTase did indeed markedly decrease the initial rate of cyclisation activity. The inhibition was non-competitive and
403 followed Hill kinetics. The inclusion of 10% (v/v) DMA had no effect on this inhibition. The reverse (coupling) reaction also lowers the yields of CD obtained: small linear dextrins which form in the reaction mixture act as acceptors for glucose residues from the CD. Inhibition of this reaction by organic co-solvents might explain the increases observed in CD production. The hydrolysis and transglycosylation of both a- and 13CD with and without solvents and with and without maltose as acceptor were followed by HPLC (Table 4). TABLE 4. Percentage of CD remaining after 4 h incubation with T-CGTase in the presence and absence of various additives (2.5 g/1 maltose or 10% v/v organic solvent) %CD remaining Additives CGTase CGTase CGTase CGTase CGTase CGTase
+ + + + +
acetonitrile dimethylacetamide maltose maltose + acetonitrile maltose + dimethylacetamide
a-CD
~-CD
76 92 69 39 74 35
92 82 95 65 65 82
In the absence of solvents or maltose, ct-CD was hydrolysed to a greater extent than 13CD and hydrolysis of both CDs was increased in the presence of maltose. The extent of hydrolysis was affected by the addition of and by the nature of co-solvent. Acetonitrile (10% v/v) reduced the extent of hydrolysis of a-CD but increased the hydrolysis of ~-CD. In contrast, addition of DMA increased hydrolysis of a-CD but decreased 13-CD hydrolysis. In the presence of maltose, acetonitrile protected et-CD against hydrolysis but had no effect on hydrolysis of 13-CD, but DMA had the reverse effect, protecting 13-CD but not ct-CD. 4. DISCUSSION. It seems likely that the effects of organic solvents on CGTase could be a combination of various factors which depend on the bacterial origin of the enzyme and the nature of the co-solvent. It is clear that the solvent effects are more subtle than can be explained simply by the log P of the solvent and it is possible that water molecules may be displaced by the solvents at specific sites either on the surface of the enzyme molecule or in the active site region. The effectiveness of low concentrations of solvents encourages speculation that solvents may interact more effectively at locations in the active site. The determination of the molecular basis of observed differences in enzyme selectivity is one of the most challenging tasks for molecular biologists. The CGTases provide a group of enzymes eminently suitable as model systems for this task. Organic solvents seem likely to be useful as additional tools in this effort.
404 References.
1. Armbruster, F.C. In: Proceedings of the 4 th International Symposium on Cyclodextrins. (Huber, O. & Szetli, J. Eds). Kluwer Academic Publishers, Dordreeht, 1988, 33-39. 2. Schmid, G., Huber, O.S. & Eberle, H. J.. lbid, 71-77. 3. Tomita, K., Tanaka, T., Fujita, Y. & Nakanishi, K. J. Ferment. Bioeng. 70 (1990) 190-192. 4. Lee, Y.D. & Kim~ H.S. Enzyme Microb. Tectmol. 13 (1991) 499-503. 5. Norman, B.E. & Jorgensen, S.T. Denpun Kagaku 39 (1992) 101-108. 6. Lawson, C.L., van Montfort, 1~, Stropotykov, B., Rozeboom, H.J., Kalk, I~H., de Vries, G.E., Penninga, D., Dijldauizen, L & Dijkstra, B.W.J. Mol. Biol. 236 (1994) 590-600. 7. Laane, C., Boeren, S., Vos, K. & Veeger, C. Bioteclmol. Bioeng. 30 (1987) 8187. 8. Bergsma, J., Bruinenberg, P.M., Hokse, H. & Meiberg, J.M.B. In: Proceedings of the 4th International Symposium on Cyclodextrins. (Huber, O. & Szejtli, J. Eds). Kluwer Academic Publishers, Dordrecht, 1988, 41-46. 9. Penninga, D., van der Veen, B.A., Knegtel, IkM.A., van Hijum, S.A.F.T., Kozeboom, H.J., Kalk, I~H., Dijkstra, B.W. & Dijkhuizen, L. J. Biol. Chem. 271 (1996) 32777-32784. 10.Knegtel, I~M.A., Strokoptykov, B., Penninga, D., Faber, O.G., Rozeboom, H.J. Kalk, K.H., Dijldmizen, L. & Dijkstra, B.W.J. Biol. Chem 270 (1995) 2925629264.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
405
Stabilization of immobilized enzymes against organic solvents: complete hydrophylization of enzymes environments by solidphase chemistry with poly-functional macromolecules. R. Fernandez-Lafuente a, C.M. Rosell a, L. Caanan-Haden b, L. Rodes b, J.M. Guisan a* a Departamento de Biocatalisis. Instituto de Catalisis. CSIC. 28049 Madrid. Spain. b Centro de Ingenieria Genetica y Biotecnologia. La Habana. Cuba.
1. INTRODUCTION The performance of enzyme biotransformations in non-conventional media is becoming increasingly popular. The use of cosolvents, non-miscible organic solvent and supercritical fluids (1-2) may be an excellent tool to shift thermodynamic equilibrium towards synthesis, to increase the solubility of reagents, to "modulate" enzyme properties, etc. However, the interaction of enzyme molecules with high concentrations of cosolvents or with hydrophobic solvent interfaces may promote their rapid inactivation. This is hindering the industrial applicability of such exciting biotransformations, when working in organic media, hydrophobic solvents may directly interact with the enzyme surfaces, promoting deleterious conformational changes. From this point of view, we may assume that a substantial increase in the "hydrophilicity" of the enzyme surface could prevent many deleterious effects of hydrophobic solvents (direct interaction and conformational changes) (3). However, the achievement of a substantial increase in the hydrophilicity of an enzyme surface via site-directed mutagenesis is not a very simple task (4). This tool for protein engineering requires the design of a large number of site-mutations on enzymes whose 3D structure need to be well known. However, the structure of many industrial enzymes has not been elucidated. In any case, the design of a large number of mutations could promote important alterations in the folding of the protein. In this point, it is worthy to note that the chemical modification of previously folded enzymes should only involve surface groups. Thus, protein engineering via chemical tools could have some advantages and it could be rationally performed on enzymes with unknown structure. However, a large number of chemical modifications on the enzyme surface could also promote dramatic reductions of the enzyme activity. "Hydrophilization" of enzymes via chemical modification of immobilized derivatives with polyfunctional macromolecules. This approach could have a number of advantages:
406
i.- Solid-phase chemical modification of proteins may be much easier than modification of soluble enzymes (5-6). A number of key features strongly facilitate the chemical modification of immobilized enzymes: intermolecular processes are now impossible, the process is very easily controlled, modification can be carried out in a wide range of experimental conditions, etc. ii.- As any other chemical modification, modification with poly-functional macromolecules is always directed towards the enzyme surface and it permit to incorporate any type of chemical moiety "on or very close to" the enzyme surface. iii.- The controlled modification with poly-functional macromolecules may promote dramatic changes in the environment of every enzyme molecule but with a minimal chemical modification of the enzyme surface. iv.- In addition to direct modification of the enzyme, the possible co-immobilization of enzymes and macromolecules on large support surfaces could permit strong modifications on the nano-environment of the enzyme molecules without any chemical modification on the enzyme structure. v.- This approach can be developed on immobilized-stabilized derivatives (e.g. multipoint covalently immobilized derivatives). In this way, the stabilization by "hydrophilization" could be additive to previous stabilization by "rigidification" associated to immobilization. Penicillin G acylase from Escherichia coil (PGA) is a very interesting enzyme with current industrial application for the selective hydrolysis of penicillin G to produce 6amino penicillanic acid (a key intermediate for the synthesis of r~-Iactam antibiotics) (7). Moreover, other interesting prospects of PGA in fine chemistry have been already reported (7, 8) synthesis of antibiotics, resolution of racemic mixtures of chiral intermediates, protection/deprotection reactions. PGA is very sensitive to the deleterious effects of organic solvents and this feature strongly limits its synthetic applicability. The chemical modification with polyfunctional macromolecules as a tool to stabilize a multipoint covalently immobilized PGA is here reported. Two different approaches are discussed: a.- the co-immobilization of the enzyme plus poly-aminated macromolecules and b.- the direct slight chemical modification of the immobilized derivatives with poly-aldehydes. Finally, the combined effect of both stabilizing approaches was also checked.
2. M E T H O D S . Enzyme activity was checked by hydrolyzing 10mM penicillin G in 100mM sodium phosphate buffer pH 8 at 37~ Penicillin G acylase was immobilized in 10% glyoxyl agarose kindly donated by Hispanagar and prepared as previously described (6). Immobilization was performed in the presence of phenylacetic acid as previously described (5,6), offering 101U per ml of support (that means less than 3% of maximum support capacity for protein immobilization). Aldehyde-dextrans (molecular weigh of 6,000 D) were prepared as described elsewhere (9). The polyethylenimine used has a molecular weight of 25,000 D. Modifications with aldehyde-dextrans were performed at 4~ and finished after 24h by adding sodium borohydride (9). Inactivations were performed at different temperatures and pH values. To initialize the experiment, the derivative was washed with 10 volumes of the desired organic solvent/aqueous buffer mixture, then suspended in 9 volumes of this mixture.
407
3.- R E S U L T S .
3.1. Co-immobilization of PGA and polyethylenimine. The co-immobilization required a three-step protocol because of the polyamine/enzyme competition by the support. Polyethylenimine is far much reactive with the support than the protein and prevents the covalent attachment of the enzyme to the support if direct competition is permitted. The experimental protocol was the following: 1.- the enzyme is immobilized for 4.5 hours. 2.- an excess of polyamine is added 3.- the suspension was reduced with sodium borohydride after 30 minutes. Reflecting the absence of chemical modification of the enzyme, PGA activity was fully preserved during the co-immobilization protocol (Table 1). However, the stability in the presence of DMF of PGA co-immobilized with polyethylenimine is almost identical to the standard derivative (Table 1).
TABLE 1.- Effect of the different treatments in the activity and stability of the immobilized PGA in the presence of 70% DMF. Treatment
Residual Activity,
(%)a
Relative Stability b
None
100
1
Polyamine/PGA Co-immobilization
100
1.05
Aldehyde -dextra n
70-75
0.85
Combined approach
77-82
50
Inactivations were performed at 4~ a Residual activity refers to the initial activity offered in the immobilization. Relative stability takes as the unit the stability of standard non-modified PGA derivative under the described conditions.
3.2. Modification of immobilized PGA with polyaldehyde-dextran. The chemical modification of the immobilized PGA caused a small reduction in the enzyme activity (25-30%). However, Table I shows that this chemical modification did not promote any significant stabilization of PGA in the presence of 70% DMF.
3.3. Modification with aldehyde-dextran of co-immobilized polyamine-PGA. The modification of the co-immobilized polyethylenimine-PGA derivative with aldehyde dextran promoted a significant stabilization of the enzyme in the presence of
408 70% DMF (a 50-fold factor) with a moderate decrease in the enzyme activity (activity decreased by around 20%) (Table 1). TABLE 2.- Effect of the combined approach in the PGA stability under different conditions
Organic solvent
Concentration
pH
T, C ~
Stabilization" factor
Dioxane
70%
7
4
300
Dioxane
80%
7
4
320
Tetraglyme
90%
7
4
>1,000
Tetraglyme
90%
8
4
>1,000
8
60
6
7
63
5
a Stabilization factor is the increment of the half-life of the modified derivative compared to the standard derivative under the described conditions. Table 2 shows that the stabilizing effects of this modification may be extrapolated to other organic solvents. In fact, these stabilizing effects increased when the apolarity and size of the organic solvent increased (e.g., maximum stabilization is detected against tetraglyme, a 1,000-fold factor). This modified derivative remained almost fully active after 3 months of incubation in 90% of tetraglyme at pH 8 and 4~ while the unmodified derivative presented a half life under two days. On the other hand, PGA thermostability is only marginally improved by this treatment (Table 2). These results seemed to reinforce the idea that the main effects of the chemical treatment performed has not been the rigidification of the enzyme structure (5,6), but the reduction in the concentration of the organic co-solvent molecules that are able to interact with the enzyme. 4. DISCUSSION
Co-immobilization of PGA with poly-amines or the chemical modification of PGA derivatives with dextrans, when separately performed, did not exert significant stabilizing effects against the effects of organic solvents. It seems that PGA has many surface areas that are highly sensitive to organic solvents. However, a substantial increase in the stability of PGA against organic solvents was achieved when both stabilizing approaches were sequentially performed. Very likely, the enzyme became completely surrounded by three hydrophilic structures: the support, the co-immobilized polyethylenimine and the chemically anchored dextrans. This combined approach for chemical modification of immobilized enzyme derivatives seemed to yield a complete hydrophilic nano-
409
environment around the external surface of the enzyme (Scheme). As a consequence of such a mild chemical modification of the enzyme, PGA preserves a high percentage of catalytic activity in spite of being much more stable against organic solvents. At first glance, this strategy for stabilization of enzymes is based in general features of enzyme structure and enzyme mechanisms of inactivation and it seems to have general applicability to the most of industrial enzymes. Perhaps some enzymes might undergo important stabilization via a single stabilizing approach. However, the performance of the combined approach is very facile and hence we propose it as a highly suitable approach for stabilization of enzymes to work in organic media.
This stabilization of PGA derivatives with regard to the deleterious effect of organic solvents does not seem due to the impossibility of the solvent to interact with the enzyme. The very open structure of the polymers surrounding the enzyme surface and the high recovery of catalytic activity suggest that only an open hydrophilic physical barrier has been created all around the enzyme molecule. Dramatic stabilization could be explained via a more simple partition effect generated by the polymeric structures. The presence of these open and highly hydrophilic polymers fully surrounding the enzyme surfaces could promote a significant reduction of the concentration of solvents in the immediate surrounding of the enzyme (e.g. by a 25 - 50 % of the concentration in the bulk solution). This reduction of apparent solvent concentration could be enough to promote dramatic stabilizing effects. The stability of enzyme derivatives may be
410
decreased by orders of magnitude when simply doubling the concentration of solvent (e.g. from 40 to 80%) (8). The combined stabilizing approach for enzyme stabilization has been performed on multipoint covalently immobilized derivatives. In this way, stabilizing factors (hydrophilization) were additive to stabilizing effects promoted by that immobilization protocol (rigidification). Thus, PGA derivatives were 107 fold more stable (against the presence of 90% of organic cosolvent (tetraglyme)) than one-point covalently immobilized derivatives could be prepared. As far as we know, any stabilization strategy (genetic, chemical and so on) have been previously reported to yield so extremely high stabilizing factors. ( 11 - 16). REFERENCES
1.-. Kasche, V. (1986) Enzyme Microb. Technol., 8, 2-16. 2.- Khmelnitsky, Y.,L., Beiova, A.B., Levashov, A.V. (1991) FEBS Lett, 284, 267-269 3.- Fangxiao, Y., Russell, A.J. (1996) Biotechnoi. Bioeng., 49, 709-716. 4.- Alvaro, G., Russell, A.J. (1991) Methods in enzymology, 202, 620-643. 5..-Femandez-Lafuente, R., Rosell, C.M., Rodriguez, V., Guisan, J.M. (1995). Enzyme Microb. Technol. 17, 517-523. 6.- Fernandez-Lafuente, R., Rosell, C.M., Alvaro, G., Guisan, J.M. (1992) .Enzyme Microb. Technoi 14, 489-495. 7.-Sheawale, J., G., Deshpande, B.S, Sudhakaran, V.K., Ambedkar (1990) Process Biochem. Int., 6, 97-103. 8.-Femandez-Lafuente, R., Rosell, C.M., Guisan, J.M.. (1996) Biotechnol. Appl. Biochem.24, 139-143. 9.-Guisan, J.M., Rodriguez, V., Rosell, C.M., Soler, G., Bastida, A., FemandezLafuente, R., Garcia-Junceda, E. (1997) in Methods in Biotechnology 1. Immobilization of enzymes and Cells (Bickerstaff, G.F. ed) Humana Press. Totowa, New Jersey. 10.-Alvaro, G., Femandez-Lafuente, R., Blanco, R.M., Guisan, J.M. (1990) Appl. Biochem. Biotechnol., 26, 181-195. 11.-Fernandez-Lafuente, R., Rosell, C.M., Piatkowska, B., Guisan, J.M. (1996) Enzyme Microb. Technol., 17, 517-523 12.-Waldman, H., Sebastian, D. (1994) Chem. Rev., 94, 911-937 13.- Favino, T.F., Fronza, G., Fuganti, C., Fuganti, D., Grasselli, P., Mele, A. (1996) J. Org. Chem., 61,8975-8979. 14.-Cardillo, G., Tolomelli, A., Tomasini, C. (1996)J, Org. Chem., 61, 8651-8654. 15.- van Straten, N.C.R., Duynstee, H..I., de Vroom, E., van der Marel, G.A., van Boom, J.H. (1997) Liebigs Ann/Recueil, 1315-1220. 16.- Margolin, A.L. (1993) Tetrahedrum Lett., 34, 1239-1242.
Stability and Stabilization of Biocatalysts A. BaUesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
411
Effect of sorbitol on immobilized a-chymotrypsin thermostability in low-water system. T. de Diego, P. Lozano, M.J. Niguez and J.L. Iborra. Department of Biochemistry and Molecular Biology "B" and Immunology. Faculty of Chemistry. University of Murcia. P.O. Box. 4021, E-30100 Murcia, Spain. The influence of sorbitol (0 to 80 mmol/g of protein) on the thermal stability of immobilized a-chymotrypsin has been studied in a low water system (hexane:ethanol:water, 69:30:1 v/v) from 30 to 60 ~ The results obtained were analyzed by a two-step series-type deactivation model, involving an active intermediate enzyme state, with excellent agreement. In all cases, the immobilized derivative exhibited non-first order deactivation kinetic, where the main protective effect of sorbitol was observed in the first-step of the deactivation profile. For all assayed temperatures, sorbitol played a clear stabilizing role only at the lowest assayed concentration (20 mmol sorbitol/g enzyme), as it was observed by both thermodynamic and kinetic analysis of deactivation profiles. However, only for the lowest assayed temperature, the increase in sorbitol concentration enhanced proportionally the etm3ane stability, obtaining a maximum increase in half-life of 40-times. The overall influence of sorbitol was related with the water activity (Aw) in the microenvironment of the enzyme. 1. INTRODUCTION Understanding parameters that control the activity and stability of enzymes in nonconventional media is essential to optimize bioeatal3r processes with industrial interest [ 1, 2] Synthetic reactions with hydrolytic enzymes, containing only a small amount of water are especially attractive (i.e. enzymatic peptide synthesis) because the equilibrium is fully directed to the synthetic way [3-5]. However, the catalytic activity of enzymes suspended in organic solvents strongly depends on the amount of water available to maintain the active conformation [6-8]. Additionally, in low water systems, the enzyme deactivation processes can be due to the direct interactions of protein with solvent molecules from, the bulk organic liquid at the phase interface, or dissolved in an aqueous phase around the enzyme [3]. Different strategies have been developed to enhance the enzyme stability in low water systems. Immobilization and/or chemical modification of enzymes to preserve the hydrophilic microenvironment of catalyst have been shown as advantageous to improve both the catalytic action and the thermostability of enzymes [6,7]. The addition of water-activity depressing agents (i.e. polyols, salts) induce favourable partitioning of available water and its retention near the enzyme, involving the increase in stability in organic media [2,9]. The aim of this paper is to analyze the thermostability of ~-chymotrypsin immobilized in hexane:ethanol:water (69:30:1 v/v) and the influence of sorbitol, as stabilizing agent, by using a series-ta/jae kinetic model. * Acknowledgements. This work was partially supported by the "Comisi6n Interministerial de Ciencia y Tecnologia", CICYT,N~BIO96-1016-C02-01.Spain.
412 2. MATERIALS AND METHODS 2.l.Materials cx-Chymotrypsin (EC 3.4.21.1) type II from bovine pancreas was purchased from Sigma Chemical Co. N-Acetyl-L-tyrosine ethyl ester (ATEE, Sigma Chemical Co.) was used as standard substrate. Celite 545 (0,01-0,04 mm particle size) was obtained from Merck. All remaining reagents were of analytical grade and used without additional purification. 2.2. Methods The immobilization method was as follows: 15 nag of a-chymotrypsin were dissolved in 7 rnL of 0.1 M phosphate buffer pH 7.8 and mixed with 1 g of oolite. The mixture was shaken for 30 rain at room temperature and then lyophilized. The immobilized derivative showed an esterase activity toward N-acetyl-L-tyrosine ethyl ester of 12.8 U/mg support and a water content of 15.4 mg of H20/g support, measured by the Karl-Fischer method [8]. The esterase activity of cx-chymotrypsin was determined by the pH-stat method using a videotritrator VIT-90 equipped with autoburette and sample station (Radiometer, Copenhagen) [8, 9]. The protocol was as follows: a 3 mL sample of 50 mlVl ATEE in 30% w/v aqueous ethanol solution containing 20 mM CaCI2 was placed into a thermostated (40 ~ reaction vessel. The reaction was started by addition of 50 lxL of suspension of immobilized derivative in organic media previously homogenized. The pH was maintained constant at 7.0 by continuous addition of 50 ~ NaOH as tritrant. One unit of activity was defined as the amount of enzyme that hydrolyzed 1 ~tmol of ATEE/min under standard conditions of assay. The stability in low-water-system was determined as follows: into a screw-capped test tube containing 1GO nag of immobilized derivative, 30 IXL aqueous sorbitol solution ( 0 - 4 M) were added. Then, the system was completed by the addition of 2.97 mL of the organic solvent (hexane:ethanol, 69:30 % v/v), and the mixture was incubated at 30, 40, 50 or 60 ~ At regular intervals of time, homogeneous aliquots of 50 lxL were extracted from the incubation mixture and the residual activity was determined immediately as described above. The results obtained were fitted to model theoretical curves using a non-linear regression program of iterative convergence by the Marquardt-Levenberg algorithm method included in the Sigmaplot 2.0 (1994) software. Water activity was determined using a humidity and temperature digital indicator HUMIDAT-IC II (Novasina, Zurich, Switzerland), with a humidity sensor model BS-3(4)/PP (Novasina). The humidity sensor was checked with control saturated salts solutions (LiCI, Aw= 0.11, Mg(NO3)2,Aw = 0.54; BaCI2, Aw = 0.91) for the overall measuring range. 3. RESULTS AND DISCUSSION The influence of temperature on the immobilized cz-chymotrypsin stability in hexane:ethanol:water system, 69:30:1% v/v has been studied between 30 and 60 ~ with and without the presence of sorbitol during 100 hours. Figure 1 shows the deactivation profiles, depicted by experimental points of immobilized cx-chymotrypsin derivative in some assayed conditions. As it can be seen, the increase in temperature enhanced clearly the loss in activity of the immobilized derivative. However, the substitution of the aqueous fraction of the medium ( 1 % v/v) by 1 M sorbitol solution in water (20 mmol/g of protein) resulted in a clear improvement of enzyme stability. This results should be explained by the "immobilization" of sorbitol in the microenvironment of the enzyme and its ability to preserve the water shell
413 around the protein molecule, which aids to maintain the essential solvophobic interactions of the native structure [2, 9]. Analysis of these data on c~-chymotrypsin deactivation by a first-order model did not represent adequately the enzyme deactivation kinetics, being noticeable their biphasic lOOq character [8 -11 ]. In this order, a series-type model of enzyme deactivation was applied 80 [10]. This model involves two first order steps, with one active intermediate (El) and a final enzyme state (E2) with possible non-zero activity, as follows, ~ 4o ,~ ...e ...... kl k2 E ~ E1 > E2 (1) 20
Of,1
(/,2
E"
0
-
2
-
4
90
95
100
where kl and k2 are first-order deactivation Time ( h ) rate constants (t1"1); and oh and 0~2 are the ratios of specific activities E1/E and E2/E, respectively. Considering as initial conditions Figure 1. Deactivation profiles of Et=o = Eo, and E1 t=o = E2 t=o = 0, the specific immobilized ct-chymotrypsin in hexane: activity of each enzyme form can be ethanol:water, 69:30:1% v/v at different determined by the integration of the rate temperatures [30 (o), 40 (11), 50 ( 0 ) and equation of each step [11]. At it was 60 ~ (A)]. Dotted line shows the experimentally proved, the final state of the deactivation profile of enzyme with 20 immobilized derivative (E2) is fully mmol sorbitol / g protein in hexane: deactivated, and the ct2 parameter of the ethanol: water (69:30:1% v/v). model should be equal to 0 [8].The fractional remaining activity a can be given by a weighted average function of specific activities of all enzyme states yields, as follows: [E + oh E1 + or2 E2] a=
= 1 + [~lkl/(k2-kl)]" { [exp(-klt)] - [exp(-k2t)] }
(2)
E0 The fit of the experimental data of Figure 1 by this equation using a non-linear regression method of iterative convergence yields the theoretical curves depicted in the same Figure. The suitability of the model was indicated by the good agreement between experimental and theoretical data. By using this model, the kinetic of the deactivation process of the immobilized derivative was analyzed at different temperatures (30 - 60~ Additionally, the influence of sorbitol concentration into the aqueous fraction of the medium (hexane:ethanol:water (69:30:1) was also studied. In all cases, kl was significant larger than k2, and k2 was practically unchanged and very close to zero. Thus, an eventual stabilization of the average enzyme activity a, at a level lower than ctl was observed. Figure 2 shows the evolution of deactivation parameters (Ctl and kl) of the immobilized derivative as a function of both the temperature and the sorbitol content (relative to the amount of protein). As it can be seen, both parameters were very
414 sensitive to the increase in temperature, increasing the k~ parameter (from 30 to 60 ~ the k~ parameter was increased in 15-times) and decreasing the oq parameter, respectively. Additionally, the presence of sorbitol not change the rate constant of activity decay (k~), showing that the deactivation process during the first step of the proposed model was dependent of the assayed temperature rather than the protective effect of sorbitol. The positive influence of sorbitol was observed by the enhancement of the o.4 specific activity of the intermediate state (czl). 0.2 However, only for 30 ~ the increase in the cz~ parameter was proportional to the sorbitol 0.0 content, while for the remaining assayed temperatures, the stabilization effect of sorbitol was only observed at the lowest assayed ~ 2 concentration. The high hydrophilicity of 1 sorbitol allows to establish that the major 0 amount of this additive should be placed into the 0 20 40 60 80 microenvironment of the immobilized enzyme. Sorbitol content Thus, the enhancement of the immobilized cz( m m o l / g protein) chymotrypsin stability by sorbitol must be explained by both the maintenance of the Figure 2. Effect of sorbitol content of the enzyme structure and the interaction with water immobilized derivative on the kinetic molecules. Gekko and Timasheff[12] concluded deactivation parameters at different tempethat the low hydrophobicity of polyols ensured ratures [30 (o), 40 (m), 50 ( 0 ) and 60 ~ that the solvatation shell around the exposed (A)]. non-polar groups remain intact at the assayed temperature, leading to protein stabilization. Furthermore, the presence of ethanol, as hydrophilic vehicle, into the organic media must give a partition of sorbitol concentration from the micro- to the macroenvironment of the immobilized enzyme. Thus, by an increase in temperature, ethanol enhanced its ability to reduce the sorbitol concentration near to the enzyme, and this fact should be related with the loss in its stabilizing function. Additionally, as Aw-reducing agent, sorbitol reduces the concentration of "free" water molecules in the microenvironment of the enzyme by increase in the hydrogen bonds interaction. Figure 3 shows the evolution of the Aw parameter of the immobilized enzyme as a function of the sorbitol content. As it can be seen, in this "dry" system, the initial Aw (0.67) is dearly reduced with respect to an aqueous medium, and the increase in sorbitol concentration produced an additional Aw reduction. So, the negative effect of sorbitol at high concentration (Figure 2) could be related with its ability to remove water molecules from the essential hydration layer of the enzyme, which could contribute to the break out of hydrophobic intramolecular interactions, to finally yield inactive protein molecule [2, 8, 9]. In this way, at all assayed temperatures, an optimum sorbitol concentration was observed (20 mmol/g protein) due to the most favourable environment.
415
0.70
<
40
0.65
"U
0.60
w .>_
0.55
2
0.501
0
,
20
,
40
,
60
80
Sorbitol content
(mmol / g protein)
Figure 3. Evolution of the Aw parameter of the immobilized enzyme suspended into the organic medium hexane:etanol:water, 69:30:1 v/v as a function of the sorbitol content at 30 ~
0
20
40
60
80
Sorbitol content (mmol/g protein)
Figure 4. Protective effect of sorbitol on immobilized o~-chymotrypsin stability at different temperatures [30 (e), 40 (11), 50 (O) and 60 ~ (A)].
In order to quantify the effect of sorbitol on the immobilized cz-chymotrypsin thermostability in this low water system, a protective effect can be defined as the ratio of enzyme half-life in the presence of sorbitol to the half-life of enzyme without sorbitol. Figure 4 shows the evolution of the protective effect of sorbitol on enzyme stability as a function of its concentration for the different assayed temperatures. As it can been seen, only at 30 ~ the protective effect of sorbitol was proportionally enhanced with its concentration. In this case, it may be seen that the protective effect obtained by using 80 mmol/g protein was near to 40. However, for all remaining temperatures, the influence of sorbitol was practically unobserved, showing a light increase in stability for the lowest assayed concentration (20 mmol/g protein). Another way to quantify the stabilization capability of sorbitol towards thermal deactivation in hexane:ethanol: water (69:30:1 v/v) is given by the thermodynamic parameters. The change in the standard free energy of denaturation for the first step A(AG~) is a good measurement of ot-chymotrypsin stabilization, which can be obtained from kinetic data and equations (3) and
(4):
AG~ = - RT In ~
k~ h KBT
(3)
A(AG~ = (AG~ (s)- (AG~ (0)
(4)
where k~ is the deactivation rate constant for the first=step (hq), KB is Boltzmann's constant (J-~ h is the Planck's constant (J.h), R is the gas constant (J-mol-l-~ T is the temperature (~ and (dG~ and (dG~ are standard free energy changes for enzyme denaturation in hexane:ethanol: water (69:30:1 v/v) with and without sorbitol, respectively.
416 Figure 5 shows the resulting values of A(AG~) at the different assayed temperatures (30-60 ~ plotted as a function of the sorbitol contem. Newly, only at 30 *C, the A(AG~) 1.2 parameter resulted in a proportional increment 0 E 0.6 with sorbitol content, showing a clear stabilizing effect, because the net free energy of protein -.- 0.0 stabilization is generally s m a l l [9,12]. Triantaf3tllou et al [2] explained that the .~ -0.6 addition of a remarkable amount of sorbitol to the solution results in the preservation of or-1.2 helix structure, a conformation less rigid than 0 20 40 60 80 the 13-sheet analyzed by the denaturation Sorbitol content temperature for various preparations of r (mmol/g protein) chymotrypsin. However, sorbitol was not able to avoid the denaturation effect of the organic medium when the temperature was increased. In Figure 5. Sorbitol content dependence of this way, at high concentration, sorbitol seems A(AG~ for immobilized ot-chymotrypsin deactivation at different temperatures [30 to be a "water-stripping" agent, enhancing the (o), 40 (m), 50 ( 0 ) and 60 ~ (A)]. enzyme deactivation process. In conclusion, sorbitol is a good stabilizing additive of the immobilized a-chymotrypsin in low-water system, where its concentration must be optimized as a function of the reaction temperature. In fact, the changes produced in the microenvironment of the immobilized enzyme by the presence of sorbitol should be considered as being an immobilization of the water molecules, rather than a rigidification of the protein structure. In this way, the Aw of the enzyme environment seems to be the key parameter in the control of enzyme stability. All results clearly showed how by changes in the microenvironment of the enzyme, the enzyme stability can be greatly improved. REFERENCES 1. F.H. Arnold, TIBTECH., 8 (1990) 244. 2. A.O. Triantafyllou, E. Wehtje, P. Adlercreutz and B. Mattiasson, Biotechnol. Bioeng., 54 (1997) 67. 3. P. Hailing, Enzyme Microb. Technol., 16 (1994) 178. 4. P. Lozano, T. de Diego and J.L. Iborra, Biotechnol. Lett., 17 (1995) 603. 5. P. Lozano, T. de Diego and J.L. lborra, Biotechnol. Lett., 19 (1997) 1005. 6. V.V. Mozhaev, Y.L. Khmelnitsky, M.V. Sergeeva, A.B. Belova, N.L. Klyachko, A.V. Levashov and K. Martinek., Eur. J. Biochem, 184 (1989), 597. 7. R.M. Blaneo, P.J. Hailing, A. Bastida, C. Cuesta and J.M. Guishn, Biotechnol. Bioeng. 39 (1992) 75. 8. P. Lozano, T. de Diego and J.L. Iborra, Biotechnol. Prog., 12 (1996) 488. 9. P. Lozano, D. Combes, and J.L. lborra, J. Biotechnol., 35 (1994) 9. 10. J.P. Henley and A. Sadana, Bioteehnol. Bioeng., 28 (1986) 1277. 11. P. Lozano, T. de Diego and J.L. Iborra, Eur. J. Biochem., 248 (1997) 80. 12. K. Gekko and S.N. Timasheff, Biochemistry, 20 (1981) 4677.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
417
Stability of immobilized enzyme-polyelectrolyte complex against irreversible inactivation by organic solvents V. Levitsky 1'2, P. Lozanol, A. Gladilin2, J.L. Iborra I 1Department of Biochemistry and Molecular Biology "B" and Immunology. Faculty of Chemistry. University of Murcia. P.O. Box 4021. E-30.100. Murcia. Spain. 2
M. V. Lomonosov Moscow State University, Chemistry Department, Ertzymology Division, 119899 B-234 Moscow, Russia
Proteolytic enzymes immobilized by adsorption on a solid support are widely used for synthetic reactions in low water systems. However, stability of such derivatives has to be optimized to avoid of reversible and irreversible inactivation of enzyme by organic solvent. Immobilization of enzyme, previously involved in non-covalent complex with polyelectrolyte, could be a possible way to suppress inactivation processes. In the present work, noncovalent complex between a-chymotrypsin and polycation Polybrene was immobilized on Celite. Stability of this biocatalyst against irreversible inactivation by organic solvents as DMSO, DMFA and AcN at 30 ~ was studied. It was observed that the complexation with Polybrene could lead to either stabilization or destabilization of enzyme in dependency on type and concentration of organic cosolvent. Maximal protective effect, defined as ratio of half-life times of enzymes, immobilized in/out of complex with Polybrene, was about 50-fold for inactivation by dry DMFA and 30-fold for inactivation by dry AcN and 20% solution of DMSO. 1. INTRODUCTION Biocatalytic systems in homogeneous water-cosolvent binary mixtures have been the subject of numerous investigations, aimed at solving both fundamental and applied problems, because these media provide a number of obvious merits [1]. However, one of the serious disadvantages of these systems is that enzymes are often inactivated by organic component [ 13]. A number of approaches, directed toward better adaptation of enzymes to water-cosolvent mixtures, have been put forward [4-7]. Formation of noncovalent ertzyme-polyelectrolyte complexes has been suggested as a new perspective approach to obtain biocatalysts with enhanced resistance against inactivation by organic solvents [8]. Protein-polyelectrolyte complexes are spontaneously formed in aqueous solutions. Multiple electrostatic interactions of counter-ionic groups of polyelectrolyte and protein surface lead to multipoint binding of a protein to a polyelectrolyte. In the presence of organic solvent this binding could be stronger * Acknowledgements. This work was partially supported by the Comisi6n Interministerial de Ciencia y Tecnologia CICYT, N~ BI096-1016-C02-01. V. Levitsky has fellowship from Ministerio de Educaci6n y Cultura, M.E.C. Spain.
418 due to the lower dielectric constant of the medium. This approach allows to obtain stabilized organosoluble biocatalytic systems for nonaqueous medium [8]. In the other hand, it is often beneficial to deposit the enzyme on a solid support. Adsorption of enzyme on a support enables to avoid aggregation, autolysis of proteolytic enzymes and to facilitate separation of the biocatalyst from the reaction mixture. It can directly affect on stability of enzymes due to immobilization procedure and direct interaction between support and enzyme. The support material can indirectly influence the catalytic activity and stability of adsorbed enzymes by affecting the partitioning of reactants and water molecules in water-organic mixture between support, enzymes and solvent [9]. In the present work we have tried to combine both updated approaches for stabilization of enzyme against inactivation by water-miscible organic solvents. Noncovalent complex between o~-chymotrypsin and positively charged polyelectrolyte Polybrene has been immobilized on Celite. Stability of this biocatalyst against irreversible inactivation by water-miscible organic solvents has been studied. 2. MATERIALS AND METHODS 2.1. Materials ot-Chymotrypsin (CT) (EC 3.4.21.1) type II from bovine pancreas was purchased from Sigma. Polybrene (PB) (hexadimethrine bromide, trademark name of 1,5-dimethyl-l.5diazaundecamethylene polymethobromide) with molecular mass 6.5 kDa was used as polycation. Celite 545 (0.01-0.04 mm particle size) was obtained from Merck. N-Acetyl-LTyrosine ethyl ester (ATEE) (Sigma) was used as a standard substrate. Dimethylsulfoxide (DMSO), N,N'-dimethylformamide (DMFA), acetonitrile (AcN) of analytical grade were obtained from Merck. 3-[N-morpholino]-propanesulfonic acid (MOPS) (Sigma) and other buffer components were of analytical grade. 2.2. Methods
The complex of CT with PB (CT-PB) was prepared by mixing of solutions of the enzyme and the polycation, both dissolved in 10 mM aqueous buffer of sodium salt of MOPS, pH 7.5. For obtaining soluble complex, the CT/PB ratio was kept equal to 1:2.8 g/g (1:10.75 mol/mol) and enzyme concentration was 0.2 mM [8]. The immobilization procedure was as follows: 3 g of Celite were suspended in 5 mL of 10 mlVI MOPS buffer, pH 7.5. After 15 rain of gentle stirring of suspension, the excess of liquid was removed by decantation and moist Celite was mixed with 9 mL of 0.2 mlVl solution of CT or CT-PB in the same buffer. After 30 rain of gentle stirring, the mixture was frozen at -30 ~ and lyophilized. The esterase activity of immobilized CT or CT-PB was determined by titration at pH 7.5 The pH 7.5 was chosen as a compromise between pH 8.0 and 6.8, corresponding to the maximal activity of CT and CT-PB, respectively [8]. A videotitrator VIT-91 equipped with autoburette ABU-91 and sample station SAIVI-90 (Radiometer, Copenhagen) was used. The protocol was as follows: a 5 mL sample of 10 ~ ATEE with 0.1 M KCI, containing a 100200 0L aliquot of previously homogenized suspension of immobilized CT or CT-PB, was placed into a thermostated at 20 ~ pH-metric cell. The concentration of organic phase in pHstate cell was not higher than 2% and has not influent the measured activity. The pH 7.5 was maintained constant by continuous addition of 10 mM NaOH as titrant.
419 The enzyme stability assay was performed as follows: into a screw-capped test tube containing 50 nag of immobilized derivative of CT or CT-PB was added 4 mL of solution of organic solvent of tested concentration in 10 mM MOPS buffer, pH 7.5. The mixture was incubated at 30 ~ with gentle stirring. At regular time intervals, the 100-200 ~tL aliquots of homogenized by stirring suspension were extracted and displaced into test tubes containing 5 mL of substrate solution with pH 3.5. After 30-min of further incubation at room temperature, the residual activity of aliquots was determined as described above. The results were fitted to theoretical model equation using a non-linear regression program of iterative convergence by the Marquardt-Levenberg algorithm (SigmaPlot 5.1).
3. RESULTS AND DISCUSSION 3.1. Protective effect of complexation with PB The inactivation of immobilized CT and CT-PB by water-miscible organic solvents was studied at 30 ~ with DMSO, DMFA and AcN at 10 concentrations ranged from 0 to 100% (v/v) For every inactivation experiment, the initial activities z~ 9 1 . . . . . . were the same and equal of the activity of aliquots of CT or CT-PB suspensions in buffer solution before incubation. As measure of protective effect 0.01 of complexation with PB, the ratio of half-life 0 20 40 60 80 100 times tla (time of lost of half initial activity) of Organic solvent, % (v/v) CT-PB and CT immobilized derivatives, was used. The protective effects observed for inactivation by different organic solvents are presented in Fig. 1. Complexation with Polybrene can lead to either stabilization or destabilization of enzyme in Figure 1. Protective effect of dependency on inactivation conditions. complexation with Polybrene, Stabilization was observed in aqueous solution measured as ratio of half-life times (tla were 50.7 h for CT-PB and 3.52 h for CT, of ot-chymotrypsin, immobilized respectively) and against inactivation by dry onto Celite in/out of complex, for DMFA (15.4 min and 0.3 min) and by dry AcN inactivation by different organic (177 h and 9.4 h, respectively). Maximal solvents: DMSO (A), DMFA (1"!) protective effect for inactivation by DMSO was and AcN (O). observed at concentration region 20-40% (at 20% ofDMSO tla were 129 h for CT-PB and 4.7 h for CT, respectively). No correlation of protective effect with medium polarity measured by the log P parameter [ 10] has been observed. To gain better understanding of observed effects, the kinetic analysis of inactivation process has been carried out. Figure 2 shows the deactivation profiles depicted by the experimental points, for inactivation of immobilized CT and CT-PB derivatives in aqueous solution (Fig. 2A) and by dry DMFA (Fig. 2B).
t
42.0
10
I
A
0.8
"~'>~o0.6 .,r
B
.....
0.4
0.2 0.0
0
10
20
30
40
50
0
2
Time (h)
4
6
8
10
Time (h)
Figure 2. Inactivation of ct-chymotrypsin (o) and its complex with Polybrene (n), both immobilized onto Celite support, at 30 ~ in aqueous media (A) and by dry DMFA (B)
The analysis of these data by a one-step first-order deactivation model did not represent adequately observed deactivation behaviour of enzyme. This result is in agreement with conclusion of different authors [11-13], that deactivation of CT follows the mechanism including irreversible lost of activity of two different in stability forms of enzyme. In this way, it was previously proposed to describe inactivation of CT, exhibiting non-first-order kinetics, by a two-step series model developed by Henley and Sadana [ 14,15] as follows:
kl E
k2 > El C~l
~ E2
(1)
C~2
where kl and 1(2 are the first-order deactivation rate constants, E, E1 and E2 are conformational states of enzyme having different specific activities, and oh and or2 are the ratios of specific activities E r E and E2/E, respectively. It was experimentally proved that the final state of CT E2 is completely irreversible deactivated conformation [ 16]. Thus, the or2 parameter of the model equal to 0. In this case the overall activity a can be written by equation 2 [ 14]. a = [1 + Otl kd(k2-k~)] [exp (-k~t)] - [cq kd(k2-k~)] [exp (-k2t)]
(2)
The computer fitting (see Methods) to equation 2 of the experimental data shown in Fig.2 yielded the values of kinetic parameter of inactivation. The theoretical curves, depicted in the same figure, were yielded by substitution all kinetic parameters in equation 2 by the calculated values. The good agreement between experimental and theoretical data (correlation coefficient higher than 0.98) showed the suitability of proposed model for describing of deactivation process of both the CT and CT-PB immobilized derivatives in water-cosolvent mixtures.
421 The influence of complexation with PB on all kinetic parameters of inactivation by DMFA is shown in Fig. 3 as evolution of ratios of values oq, k] and k2 calculated for CT, immobilized in/out of complex with PB. From the comparison of general protective effect (Fig. 1A) with contribution of each kinetic parameter (Fig. 3), the following observations can be made. The protective effect of PB in water solution can be related to increasing of relative activity oq of intermediate form E1 of CT-PB (0.8 vs 0.35 for CT) (Fig. 3A) and limitation of inactivation with rate constant k2 (Fig. 3C) from 0.043 h "1 to 0.01 hq, while stabilization action of PB on the first inactivation step was negligible (Fig. 3B). At intermediate concentrations of DMFA (low 40%), general protective effect decreased and changed to destabilization mainly due to decrease in relative activity of intermediate form (cq decreased to 0.026 for CT-PB and only to 0.33 for CT), while influence of PB on both inactivation rate constants was still slightly stabilizing. At DMFA concentration range from 40 to 60%, complexation with PB accelerated both inactivation step, but relative activity of E~ started to increase. At DMFA concentration up to 60%, the ratio of or1 values reached the constant level (0.14 for CT-PB and 0.07 for CT) and protective effect increased due to decrease of ratios for both rate constants of inactivation. It is interesting to note, that ratios of both constants k~ and k2 were changed by DMFA concentration in the same manner. This parallel change may indicate the same mechanism of PB action on both inactivation steps, and therefore, the same nature of inactivation processes for E and E~ enzyme forms
O
.p.,4
10
A
0.1 o
,I,,~
10
B
1 0.1 0.01 o 10 ".,~, 1
C
0.1 0.01 0 20 40 60 8 0 1 0 0 D M F A , % (v/v)
Figure 3. Effect of complexation with Polybrene, measured as ratios of kinetic parameters or1, kl and k2, for inactivation by DMFA of ot-chymotrypsin, immobilized onto Celite in/out of complex.
3.2. Possible reasons of stabilization and destabilization of CT by complexation with PB
Stabilization of enzyme by complexation with PB in water solution could be related to restriction in conformational flexibility due to multipoint electrostatic interactions surface charged groups of CT with charged groups of PB. This additional immobilization could result in limitation of irreversible conformational changes, presumably leading to activity lost of both enzyme forms E and El. Protective effect at high concentrations of DMFA (80-100%) could be interpreted as follows. Inactivation of enzyme in this region of solvent concentrations can be caused direct interaction of solvent molecules with protein surface, resulting in disruption of hydration shell around the protein molecule. It could contribute to enzyme desorption from the support with later aggregation as well as to the break out of hydrophobic intramolecular interactions [2, 9, 11]. During the complexion with PB, the overall concentration of charged groups in the vicinity of enzyme was increased, because PB molecule has 35 positively charged
422 quaternary amino groups and only the minority of that was taken out by interaction with enzyme charged groups [8]. These additional charged groups can retain the water molecules and preserve water shell around enzyme [2, 8]. The influence of complexation with PB on inactivation kinetics at intermediate concentration of DMFA is difficult to interpret within the frame of used simple model. Nonmonotonic change of all kinetic parameter ratios indicates more complex conformational behaviour of enzyme. The model used, however, can describe more complex cases of inactivation, including additional reversible steps, which can not be determined kineticaUy. In these cases, the parameters ct~, k~ and k2 would be global kinetic parameters, i.e. combinations of individual rate and equilibrium constants [15]. In this way, the complex mode of PB influence may be related to the influence on conformational equilibrium between forms of enzymes having different activity or/and stability. 4. CONCLUSION The nature of enzyme inactivation by organic solvents at different concentration region is still not completely clear, and in this turn, molecular mechanism of stabilization and destabilization of enzyme by complexation with polyelectrolyte has to be determined. However, the obtained results demonstrated significant potentiality of this stabilizing approach. Due to technical simplicity and universality of proposed approach, it seems to be promising for creation of high-stable biocatalysts for peptide synthesis in nearly-dry organic solvent systems. REFERENCES
1. V.L. Khmelnitsky, V.V. Mozhaev, A.B. Belova, M.V. Sergeeva, K. Martinek, Eur. J. Biochem., 198 ( 1991) 31. 2. V.V. Mozhaev, Y.L. Khmelnitsky, M.V. Sergeeva, A.B. Belova, N.L. Klyachko, A.V.Levashov, K.Martinek, Eur. J. Biochem., 184 (1989) 597. 3. Y. Tomiuchi, T. Kijima, H. Kise. Bull. Chem. Soc. Jpn., 66 (1993) 1176. 4. J.S. Dordick, Biotechnol. Prog., 8 (1992) 259. 5. Y.L. Khmelnitsky, A.V. Levashov, N.L. Klyachko, K. Martinek, Enzyme Microbiol. Technol., 10 (1988) 710. 6. V.V. Mozhaev, M.V. Sergeeva, A.B. Belova, Y.L. Khmelnitsky, Biotechnol. Bioeng., 35 (1990) 653. 7. C.-H. Wong, Trends Biotechnol. 10 (1992) 378. 8. E.V. Kudryashova, A.K. Gladilin, A.V. Vakurov, F. Heitz, A.V. Levashov, V.V. Mozhaev, Biotechnol. Bioeng., 35 (1990) 653. 9. M. Reslow, P. Adlercreutz, B. Matiasson, Eur. J. Biochem., 172 (1988) 573 10. C .Laane, J. Tramper, M.D. Lilly (ed.) Biocatalysis in organic media, Elsevier Science Publ., Amsterdam, The Netherlands, 1987. 11. V.V. Mozhaev, N.S. Melik-Nubarov, V.Y. Levitsky, V. Siksnis, K. Martinek, Biotechnol. Bioeng., 173 (1992) 147. 12. R.K. Owusu, N. Berthalon, Food Chem., 48 (1993) 231. 13. V.Y. Levitsky, N.S. Melik-Nubarov, V.A.Siksnis, V.Y. Cn'inberg, T.V. Burova, A.V. Levashov, V.V. Mozhaev, Fur. J. Biochem., 219 (1994) 219. 14. J.P. Henley, A. Sadana, Enzyme Microb. Technol., 7 (1985) 50. 15. J.P. Henley, A. Sadana, Biotechnol. Bioeng., 28 (1986) 1277. 16. P. Lozano, T. De Diego, J.L. Iborra, Biotecnol. Prog., 12 (1996) 488.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
423
E n h a n c e m e n t o f invertase activity in organic m e d i a for o l i g o s a c c h a r i d e synthesis S. Bielecki and R. I. Somiari Institute of Technical Biochemistry, Technical University of Lodz Stefanowskiego 4/10, 90-924 Lodz, Poland 1. ABSTRACT 13-Fructofuranosidase catalysed conversion of sucrose in media containing organic cosolvent and sodium dodecyl sulphate (SDS) were studied. The addition of up to 90 mM SDS resulted in an improvement in hydrolytic and transfructosylating activity of invertase averaging >65% and >98%, respectively, in media containing 60% (v/v) ethyl acetate, butyl acetate, hexane or benzene. In general, oligosaccharide synthesis was best in the presence of butyl acetate with a 4-fold increase in yield occuring following the addition of 90 mM SDS. Interestingly, reversed micelles were not formed in similar organic media containing even up to 120 mM SDS, an indication that the enhanced activity recorded in the presence of this surfactant was due to mechanisms other than entrapment in reversed micelles. Because 90mM SDS had no apparent effect on activity in aqueous medium, but did so in organic media, it is clear from these results that in spite of the absence of reversed micelles, SDS still offers some level of protection with regards to inactivation of invertase by the four organic solvents examined. 2. INTRODUCTION Invertase (13-fructofuranosidase, EC 3.2.1.26) is easily obtainable and exhibits transferase activity so can be exploited for synthesis of fructooligosaccharides using sucrose as the donor and acceptor of the transferred glycoside. But the transferase activity of this enzyme in aqueous media is typically low because water is the preferred acceptor [1]. Although it is known that the transglycosylating activity of glycosyl hydrolase may be enhanced by using high substrate concentration and/or low water media, it is not always feasible to use high substrate concentration because of the problem of solubility or substrate inhibition. With respect to invertase catalysis high substrate concentration do not correspondingly increase oligosaccharide formation. The amount of oligosaccharide synthesized by this enzyme in aqueous medium is below 10 % (w/w) even in media containing >60 % (w/v) sucrose. [2,3] It is known that adding an organic co-solvent may shift the equilibrium of hydrolase catalysed reactions towards synthesis [4]. This approach is however limited because sugar
424 substrates are not soluble in those organic solvents that are more suitable for biocatalysis [4,5]. If however, the enzyme is protected from the bulk organic phase, then the operational performance of the enzyme may be maintained. Invertase activity has been stabilized and even enhanced in media containing some water-immiscible organic solvents by entrapment in the reversed micelles formed by surfactants such as sodium lauryl sulphate [6]. But the effect of this surfactant on transfructosylation activity of invertase and hence oligosaccharide accumulation is not known. Moreover because, the hydrophobicity index of invertases obtained from different strains of the same species differ remarkably [7] similar media conditions may affect invertases differently. We recently reported on the performance of invertase in organic media containing SDS as well as some properties of the main oligosaccharide formed [8] and hereby elaborate further on the posible role of SDS in enhancing the transferase activity of invertase in organic media optimised for fructooligosaccharide synthesis. 3. MATERIALS AND METHODS
A commercial invertase (Novo Nordisk) was partially purified by ultrafiltration as previously described [3]. The activity of the preparation was determined by measuring the amount of glucose released from 0.23 M sucrose solution with 50 mM acetate buffer (pH 5) and incubated at 50~ for 15 min. After inactivation of the enzyme the glucose concentration was determined in appropriately diluted samples using a glucose oxidase test kit (Sigma). One unit of invertase activity was defined as the amount of enzyme that releases 1~tmol glucose min l under above conditions All the reactions contained approximately 150U of enzyme. Unless indicated, all reactants were reconstituted in 50 mM phosphate buffer, pH 7.5 and after mixing, vortexed for 30s before incubation. For hydrolysis, glucose released was measured enzymatically in reactions containing 0.23 M sucrose, 0-60% (v/v) ethyl acetate, butyl acetate, benzene or hexane and 0-120 mM SDS. At the end of incubation (50~ 60min) the reaction mixture was mixed vigorously and 1 part was transferred to 100 parts of HPLC grade water, before inactivation of the enzyme and quantifying the glucose released. Transglycosylation reactions contained 1.7 M sucrose, 60% (v/v) organic solvent, 90 mM SDS and enzyme. Controls without SDS and without organic solvents were set-up simultaneously. Incubation was at 50~ for 24h and the reacted mixture was treated as described above. For quantitative analysis, organic solvents were remove in a vacuum rotary evaporator at 40~ after which proteins and impurities were removed by solid phase extraction using BAKERBOND diol columns and protocol. Appropriately diluted samples were then analyzed by HPLC with refractive index detector [9]. The homogeneity of the media containing butyl acetate as bulk phase was examined by fluorescence spectroscopy in the presence and absence of SDS. About 150U of invertase was added to each set-up. After mixing and allowing the phases to separate, the steady-state fluorescence intensity of the organic phasewas determined using an excitation wavelength of 280nm at a resolution of lnm over a measurement range of 290-500nm. These spectra obtained were corrected for background values due to SDS, solvent and reconstituting buffer.
425 4. RESULTS AND DISCUSSION
4.1. Invertase activity in organic media The inversion of sucrose by 13-fructofuranosidase was affected by addition of water immiscible solvents with different logP. As presented in Figl, invertase activity ranged from 31% (ethyl acetate) to 56% (hexane) in the presence of organic co-solvents. The differences is most likely due to the direct effect of the solvent on media environment and hence the operational performance of the enzyme. It is generally believed that solvents with logP >2 cause less inactivation of enzyme. Solvent that partition in the aqueous phase can change the conformation of an enzyme molecule by interacting with essential water layer around enzyme. This generally cause greater changes to the catalytic efficiency of the enzyme. This may explain why sucrose hydrolysis was remarkably higher in hexane (logP 3,5) as compared to ethyl acatate (logP 0.68).The addition of 90 mM SDS did not produce any remarkable change in the hydrolytic activity of invertase in aqueous media. On the other hand the activity increased from 44% (ethyl acetate) to 98% (hexane) following the addition of 90 mM SDS. The percentage of sucrose hydrolysed in organic media increased with increase in SDS concentration (Fig 2), showing that the presence of SDS improves the catalytic efficiency of invertase in these organic media. This finding is consistent with the report on invertase activity in hexane, benzene and carbon tetrachloride, where reverse micelles were formed in the presence of SDS [6]. Our system was not appropriate for reverse micelles formation so it is apparent that the stabilization of invertase in these media was not due to entrapment of the enzyme in the reverse media.
Figure 1. Hydrolytic activity of invertase in aqueous media and in media containing 60% (v/v) of organic solvent. Substrate was 0.23 M sucrose prepared with 50 mM acetate buffer, pH 5.0 and incubation was at 50~ for 60min. Glucose released was determined enzymatically.
426 120 .~
o~ 100
~.
~
~.
~
~.
I
I
50
60
70
8060 o 0
40
,
20"
-
I
10
20
30
40
80
90
I
I
100
110
120
SDS [mM] Figure 2 Effect of SDS concentration on % sucrose hydrolysed by invertase in aqueous medium (O) and in media with 60% (v/v) hexane (x), benzene (A), butyl acetate (0) or ethyl acetate (71). Reaction conditions and analysis were as described for Fig. 1. 4.2. Oiigosaccharide synthesis The amount of oligosaccharides synthesized in aqueous media in the absence and presence of 90 mM SDS were .--1.6 and 1.5% (w/w), respectively (Fig.3). Thus, as observed for hydrolytic activity, this surfactant has no effect on transglycosylation activity in aqueous medium. On the other hand, oligosaccharide synthesis in organic media increased by an average of 98% when 90 mM SDS was added (Fig.3), indicating that it plays a positive role in the catalytic efficiency of invertase in the presence of organic solvents. Transglycosylating activity was best in the medium with butyl acetate, and oligosaccharide synthesis increased from 3% to 12% when SDS was added. The finding that butyl acetate is the best solvent among these four has been previously reported suggesting that the inherent attributes of butyl acetate particularly favours transglycosylation by this invertase [9]. Oligosaccharides synthesised in butyl acetate were resolved into three groups by HPLC (Table 1), and the transferred glycoside was shown to be fructose by enzyme hydrolysis and HPLC analysis (results not presented).
Table 1 Composition of oligosaccharides synthesised by invertase in media containing butyl acetate Reaction mixture* without SDS with 90raM SDS
2DP @ 6,5 4,1
Oligosaccharide composition (%)# 3DP 4DP 64,5 29,0 65,3 . . . . . 30,6
* Reaction mixture contained 1.7M sucrose, 60% (v/v) butyl acetate, and 150U invertase, pH 7.5. Incubation was at 50~ for 24h and analysis was by HPLC. § % of total. @ DP, degree of polymerisation.
427
Figure 30ligosaccharides synthesised by invertase in aqueous media and in media containing 60% (v/v) ethyl acetate, butyl acetate, benzene or hexane. Substrate was 1.7 M sucrose, incubation was at 50~ for 24h and analysis was by HPLC.
4.3. Role of SDS It was of interest to note that reversed micelles were not formed in any of the organic systems following SDS addition. Thus, the enhancement of invertase activity in these organic media was not due to entrapment in reversed micelles, as was the case in a literature report [6]. Entrapment in reverse micelles protects biocatalysts from inactivation especially by hydrophilic solvents, because such solvents have greater tendency to moderate the water concentration around the enzyme. However, based on the difference observed in the derived amino sequence and hydrophaty plots of invertase from different strains of the same species [7], it is obvious that the degree of inactivation of invertase by an organic solvent will in part depend on the hydrophobieity index of the enzyme. In any ease, the fact that enhanced activity was still recorded in the absence of reverse mieelles may be because this invertase is relatively more stable in organic media. However, using the tryptophan residue of invertase as probe, the fluorescence properties of a medium containing butyl acetate as bulk phase showed an increase in the fluorescence intensity of the organic phase when SDS was added (Fig.4). This means that SDS improved the partitioning of the enzyme as well as other reactants in the organic phase. It is known that when organic solvent/surfactant/aqueous phase ratio is not appropriate for reversed micelles formation, surfactants may still promote the formation of bi-continuous systems or lamella phases, which leads to an increase in interfacial area [10]. Results of this study suggest that this phenomenon and the stabilization effect of SDS contributed to the enhanced operational performance of invertase in the organic media examined.
428 30 A 25 20 15 I=I
o
10
~
-5 290
330
....
I'
370
1
410
I ' 450
', 490
Wavelength [nm] Figure 4 Fluorescence spectra showing the effect of SDS addition on fluorescence intensities. Spectrum A, aqueous suspension of invertase, B, butyl acetate phase after addition of 90 mM SDS and C, butyl acetate phase in the absence of SDS. Acknowledgement: Prof. S. Wysocki, Institute of General Food Chemistry, Technical University of Lodz is acknowledged for the fluorescence spectroscopy studies. REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.
P.S.J. Cheetham, A.J. Hacking and M. Vlitos, Enzyme Microb.Technol.,11 (1989) 212 A.J.J. Straathof, A.P.G. Kieboom and H. van Bekkum, Carbohydr. Res. 146 (1986 154. R.I. Somiari and S. Bielecki, Biotechnol. Lea., 17 (1995) 519. J.S. Dordick, Enzyme Microb. Technol., 11 (1989) 194. E.N. Vulfson, R. Patel, J.E. Beecher, A.T. Andrews and B.A. Laws, Enzyme Microb. Technol., 12 (1990) 209 D.B. Madamwar, J.P. Bhatt, R.M. Ray and R.C. Srivastava, Enzyme Microb. Technol. 10 (1988) 302. R.I. Somiari, H. Brzeski, R. Tate, S. Bielecki and J. Polak, Biotechnol. Lett. 19 (1997) 1243. S. Bielecki and R.I. Somiari, Biotechnol. Lett., 20 (1998) 287. S. Bielecki and R.I. Somiari, Bioeatal. Biotransf. 13 (1996) 217. P. Adlercreutz, Biocatal. Biotransf. 14 (1996) 1.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 1998 Elsevier Science B.V.
429
Effects of c r o w n ethers on the activity of e n z y m e s in peptide f o r m a t i o n in organic m e d i a ~ Dirk-Jan van Unen, Johan F.J. Engbersen and David N. Reinhoudt Laboratory of Supramolecular Chemistry and Technology University of Twente, P.O.Box 217, 7500 AE Enschede, The Netherlands Phone +31 53 4892980, Fax +31 53 4894645, E-mail [email protected]
The effects of crown ethers on the rate of protease-catalyzed dipeptide formation using suspensions of oc-chymotrypsin and cross-linked crystals of subtilisin Carlsberg in organic solvents is described. Lyophilisation of cz-chymotrypsin in the presence of 50 equivalents of 18-crown-6 results in a 450 times increased enzymatic activity in acetonitrile. Drying of crosslinked crystalline subtilisin Carlsberg from acetonitrile containing 50 mM 18-crown-6 enhances the activity 13 times. A possible mechanism for the crown ether activation is discussed.
1. INTRODUCTION Nowadays the applicability of enzymes in synthetic organic chemistry is well recognized. Investigations in the field of enzyme-catalyzed organic synthesis were further boosted by the invention that enzymes can function in organic solvents [1]. The use of enzymes in nonaqueous media offer a number of advantages over the use of enzymes in aqueous solutions, like enhanced thermal stability of the enzyme, easy regeneration of the suspended enzyme by filtration, increased substrate solubility, favorable equilibrium shift to synthesis over hydrolysis, and altered selectivity properties of the enzyme [2]. The most important drawback of the use of enzymes in organic media is the reduced activity compared to aqueous conditions. Typically, this reduction in activity is 2 to 6 orders-ofmagnitude. Optimization of the enzymatic activity in organic solvents is therefore one of the major themes of investigation in this field. Modification of charged amino acid residues on the enzymes surface by means of protein engineering or chemical modification is one of the # This investigation was supported by the Netherlands Technology Foundation (STW) and Technical Science Branch of the Netherlands Organization for Advanced and Pure Research (NWO). Dr. I. Sakodinskaya is acknowledged for valuable discussions.
430 strategies. Furthermore, immobilization and medium engineering were shown to result in increased enzyme performance. We have reported that addition of crown ethers enhances the activity of proteases [3] and tyrosinases [4] in organic solvents. High accelerations, typically in the range of 500 times, were found when the crown ether was added prior to lyophilisation [5]. Thiacrown ethers were also reported to change the enantioselectivity of lipases [6]. The low enzymatic activity in organic solvents is one of the major reasons why applications on a larger scale are still a future prospective. A reaction which is potentially very suitable for application of enzymes in organic solvents is the enzymatic synthesis of peptide bonds. The formation of peptide bonds catalyzed by proteases offers clear advantages over chemical methods, such as mild reaction conditions, no racemization, and the fact that time-consuming protection and deprotection steps render superfluous. It is obvious that in order to overcome hydrolysis problems this reaction needs to be performed in non-aqueous systems. Here, we report a study of the protease-catalyzed formation of N-acetyl-L-phenylalanyl-Lphenylalaninamide from the 2-chloroethyl ester of N-acetyl-L-phenylalanine and Lphenylalaninamide and the effects of crown ethers on this synthesis. The effects are studied using suspensions of tx-chymotrypsin and subtilisin Carlsberg enzyme crystals.
2. MATERIALS AND METHODS tx-Chymotrypsin (E.C. 3.4.21.1) type 11 from Bovine Pancreas (54 U/mg protein; hydrolysis of N-benzoyl-L-tyrosine ethyl ester at pH 7.8) and the peptide precursors N-Ac-L-Phe and LPhe-NH2 were obtained from Sigma (St. Louis, MO). Cross-linked crystals of subtilisin Carlsberg were prepared according to the method of Schmitke et al. [7]. The 2chloroethylester of N-Ac-L-Phe was synthesized from N-Ac-L-Phe and 2-chloroethanol using Amberlite IR-120 as a catalyst. 18-Crown-6 was a gift from Shell Laboratories (Amsterdam, The Netherlands). The solvents were of analytical grade or higher and were from Acros (Geel, Belgium).
2.1. Pretreatment of o~.chymotrypsin a-Chymotrypsin (5 mg/ml) was dissolved in 20 mM sodium phosphate buffer pH 7.8 containing the appropriate amount of 18-crown-6. The equivalents of added crown ether are relative to the moles of enzyme. The samples were lyophilized, after rapid freezing in liquid nitrogen, for 24 hours. For comparison studies D-sorbitol (5 mg/ml) was used instead of crown ether, using the similar procedure. 2.2. Pretreatment of cross-linked crystalline subtilisin Carisberg Cross-linked crystals of subtilisin Carlsberg (1.0 mg/ml) were soaked in acetonitrile containing the indicated concentration of 18-crown-6. Subsequently the solvent was left to evaporate overnight at room temperature.
431 2.3. Studies on the effect of 18-crown-6 on the rate of dipeptide formation All enzyme preparations, peptide precursors and solvents were equilibrated at a thermodynamic water activity of 0.113 above a saturated LiC1 solution for 24 hours. Reactions were performed in duplicate on a 1 ml scale with magnetic stirring at 500 rev./min. Typical conditions: 2.5 mg/ml r or 0.5 mg/ml subtilisin Carlsberg enzyme crystals, 50 mM N-Ac-L-Phe-OEtC1 and 50 mM L-Phe-NH2 in acetonitrile at 30~ The reactions were terminated by the addition of 4 volumes of dimethylsulphoxide. The reaction mixture was analyzed by HPLC. Initial rates were calculated from conversions <5%.
3. RESULTS AND DISCUSSION The reaction studied was the protease-catalyzed peptide bond formation between the 2chloroethylester of N-acetyl-L-phenylalanine and L-phenylalaninamide, using (x-chymotrypsin and cross-linked crystalline subtilisin Carlsberg (Scheme 1).
~
~
+
O
'~N
O
C!
N-Ac-L-Phe-OEtC1
~ H2N
protease
0
,,Jr,.
0
O
L-Phe-NH 2
N-Ac-L-Phe-L-Phe-NH 2
Scheme 1. Protease-catalyzed peptide bond formation. 3.1. Effects of 18-crown-6 on peptide bond formation using o~-chymotrypsin Initially, the catalytic activity of three proteases, namely subtilisin Carlsberg, trypsin and r chymotrypsin, present as suspensions in acetonitrile was investigated for the reaction depicted in Scheme 1. All three proteases could be activated by lyophilisation in the presence of 18crown-6. The highest acceleration in the rate of peptide bond formation was found in the case of r Therefore, further studies were performed using this enzyme. In order to optimize the activation effect, the influence of the amount of 18-crown-6 was studied (Table 1). In the presence of 50 to 100 equivalents of crown ether a maximum activity enhancement of 450 times is observed. This results in an enzymatic activity of 0.7 U/mg enzyme. Upon addition of larger amounts of crown ether the enzyme activation is decreasing. The activation effect is due to the macrocyclic nature of the crown ether while pentaglyme, the linear chain analog of 18-crown-6, has no effect on the enzyme activity. Comparative studies using D-sorbitol, a well-known lyoprotectant, instead of crown ether revealed that the
432 activation effect of 18-crown-6 was much more effective as pretreatment with D-sorbitol resulted in an acceleration of only 8 times. Table 1 Crown ether induced enhancement (V0(18-crown-6)/V0) of the ct-chymotrypsin-catalyzed peptide bond formation between N-Ac-L-Phe-OEtC1 and L-Phe-NH2 in acetonitrile. For details, see methods. Equivalents of 18-crown-6
V0
Acceleration
0 50 100 250 500 1000 2500
(nmol/min*mg protein ) 1.5 674 700 462 130 12.4 0.8
436 450 298 84 8 0.5
A hypothesis for the mechanism of the crown ether acceleration effect is that 18-crown-6 is complexing the e-ammonium functions of the lysine residues on the exterior of the enzyme. In this way protonation of the amine functions is stabilized. Also the formation of inter- and intramolecular salt-bridges between anionic sites on the enzyme and these ammonium functions in organic solvents is prevented. Formation of these salt-bridges may result in enzyme molecules with altered catalytic properties compared to the native enzyme. The most direct proof that the ammonium functions play a role in the crown ether acceleration is the observation that acetylated trypsin, in which the lysine residues have been acetylated, are not activated by crown ether pretreatment, while normal trypsin is accelerated very effectively [5]. When larger amounts of crown ethers are added one can imagine that also polar residues from the interior of the enzyme are complexed, resulting in a deformation of the native enzyme conformation and a lower catalytic activity. Conformational changes of the enzyme during lyophilisation are known to occur. Upon suspending the lyophilized enzyme powder into an organic solvent the enzyme can be locked in a distorted conformation due to lack of conformational freedom. This may explain the observed decrease in enzyme activaty at larger amounts of 18-crown-6. The crown ether induced enhancement of enzyme activity also turned out to be applicable for peptide bond formation using a variety of peptide precursors, like N-acetyl-L-tyrosine ethylester, D-phenylalaninamide, L-leucinamide, and L-tyrosinamide, and in a whole range of solvents, ranging in polarity from dioxane to toluene.
433 3.2. Effects of 18-crown-6 on peptide bond formation using cross-linked enzyme crystals of subtilisin Carlsberg An emerging technology in the field of non-aqueous enzymology is the use of cross-linked crystalline enzymes [8,9]. These mechanically and thermally stable class of biocatalysts were investigated on the effects of crown ethers. Addition of 18-crown-6 to the reaction mixture without pretreatment of the subtilisin Carlsberg crystals did not result in any acceleration. The initial peptide formation activity remained 0.44 nmol/min*mg enzyme. As freeze drying of enzyme crystals from aqueous buffer results in a distortion of the enzyme crystal lattice, due to crystallisation of the water during freezing, lyophilisation in the presence of crown ethers, as was performed for the r preparations in the previous section, is not expected to be a suitable procedure in this case. Therefore, another way of pretreatment with crown ethers has been investigated, that is soaking and drying of cross-linked crystalline subtilisin Carlsberg from an organic solvent, typically acetonitrile, in which 18-crown-6 is dissolved.
14 "N
12 10
<
/
8
0
J
i0
/
Y
z0
3o
.io
5o
[ 18-crown-6], (raM) Figure 1. Effect of the amount of 18-crown-6 on the peptide bond formation between N-AcL-Phe-OEtC1 and L-Phe-NH2 in acetonitrile using soaked cross-linked crystalline subtilisin Carlsberg. For details, see methods. Figure 1 showes that soaking and drying of the enzyme crystals results in a enhancement of the enzymatic activity. An acceleration of 13 times can be achieved by drying the enzyme crystals in the presence of 50 mM 18-crown-6. Drying of cross-linked crystalline enzymes in the presence of surfactants was also reported [9] to enhance enzymatic activity. However, most surfactants lost their activating effect after storage for a few days. In contrast, the crown ether activation turned out to be very stable as the activated cross-linked crystals had still the same activity after one month of storage at 4~ In conclusion, these results show that lyophilisation of t~-chymotrypsin and soaking of crosslinked crystalline subtilisin Carlsberg with 18-crown-6 results in enzyme preparations with
434 strongly enhanced catalytic properties in the peptide bond formation in organic solvents. Since crown ethers are readily available and the pretreatment procedure is easily performed this approach is a versatile way to overcome the usually observed large decrease in activity of enzymes in organic media.
REFERENCES
1. J.S. Dordick, Enzyme Microb. Technol., 11 (1989) 194. 2. A.M.P. Koskinen and A.M. Klibanov (eds.), Enzymatic reactions in organic media, Blackie academic & professional, Glasgow, 1996. 3. D.N. Reinhoudt, A.M. Eendebak, W.F. Nijenhuis, W. Verboom, M. Kloosterman and H.E. Schoemaker, J. Chem. Soc., Chem. Commun., (1989) 399; J. Broos, M.N. Martin, I. Rouwenhorst, W. Verboom and D.N. Reinhoudt, Recl. Trav. Chim. Pays-Bas, 110 (1991) 222; J.F.J. Engbersen, J. Broos, W. Verboom and D.N. Reinhoudt, Pure & Appl. Chem., 68 (1996) 2171. 4. J. Broos, R. Arends, G.B. van Dijk, W. Verboom., J.F.J. Engbersen and D.N. Reinhoudt, J. Chem. Soc., Perkin Trans. 1, (1996) 1415. 5. J. Broos, I.K. Sakodinskaya, J.F.J. Engbersen, W. Verboom and D.N. Reinhoudt, J. Chem. Soc., Chem. Commun., (1995) 255. 6. T. Itoh, Y. Takagi, T. Murakami, Y. Hiyama and H. Tsukube, J. Org. Chem., 61 (1996) 2158; Y. Takagi, J. Teramoto, H. Kihara, T. Itoh and H. Tsukube, Tetrahedron Lett., 37 (1996) 4991. 7. J.L. Schmitke, C.R. Wescott and A.M. Klibanov, J. Am. Chem. Soc., 118 (1996) 3360. 8. N.L. St. Clair and M.A. Navia, J. Am. Chem. Soc., 114 (1992) 7314. 9. N. Khalaf, C.P. Govardhan, J.J. Lalonde, R.A. Persichetti, Y.-F. Wang and A.L. Margolin, J. Am. Chem. Soc., 118 (1996) 5494.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
R e c o v e r y o f the transesterification
activity o f an i m m o b i l i z e d
435
lipase
after its use
in fat
Ferreira-Dias, S.a; Duarte, C.S?; Falaschi, V.b; Marques, S.R.d; Gusm~o, J.H?; da Fonseca, M.M.R. d ' Instimto Superior de Agronomia, Tapada da Ajuda, 1399 Lisboa-Codex; Portugal b Permanent address: Sez. Microb. Ind., DISTAM, Univ. degli Studi di Milano, Italy cFIMA, Produtos Alimentares, Lda., SP Iria de Az6ia; Portugal d Instituto Superior T6cnico, Centro de Engenharia Biol6gica e Quimica, Av. Rovisco Pais, 1096 Lisboa-Codex, Portugal.
1. INTRODUCTION Natural fats present an organized fatty acid distribution in their molecules which determines their functional, nutritional and organoleptic properties.. By transesterification it is possible to modify this distribution by rearranging the acyl groups among triglycerides. The changes introduced by acyl exchange may lead to an increase in the plastic range of the transesterified fat, as compared to the starting blend. Currently transesterification is carried out at high temperature (higher than 250~ without catalyst or with an acid, alkaline or metal catalyst (e.g., sodium methoxide or sodium methylate), under reduced pressure at lower temperature (70-150~ The interchange of acyl groups occurs at random and the composition reached at equilibrium obeys the laws of probability. The final products may remain contaminated by residual catalyst. The lack of specificity of chemical catalysts may lead to the formation of considerable amounts of side products (soaps as sodium salts of fatty acids, mono- and diglycerides) with a subsequent decrease in yield ~. Oils and fats have to be completely dried before transesterification, since chemical catalysts are explosive if allowed to contact water. In addition, the presence of even small amounts of free fatty acids (FFA), moisture and unsaponifiable matter causes inactivation of catalysts i. In the margarine industry, the efficency of the transesterification reaction is indirectly evaluated by the decrease in the extent of fat eristalization at a given temperature. This may be measured by dilatometry (Solid Fat Index- SFI) or by Nuclear Magnetic Resonance (Solid Fat Content- SFC) 2. The SFC values at 10~ 20~ and 35~ (SFC~0.o SFC~0.c and SFC35.c) are related to the rheological behaviour of fats at storage, packaging and consumption temperatures, respectively. Therefore, after transesterifieation, SFC35.c should be smaller than their original counterparts, to prevent a sandy and coarse texture of the margarine.
436 The use of lipases (acylglycerol acylhydrolases, EC 3.1.1.3.) in transesterification reactions has become a challenge for the oleochemical industry. The positional and fatty acid selectivity of lipases enables the production of structured TG, impossible to obtain by chemical routes ~ since the use of 1,3-selective lipases preserves the original 2-position of unsaturated fatty acids in triglycerides. The major objective of this study was to produce fat bases for the margarine industry (MFB), by transesterification of palm oil stearin (POS) with palm kernel oil (PKO), in a continuous reactor, catalysed by an immobilized commercial lipase. Since PKO is more expensive than POS, the main purpose was to obtain a suitable MFB using a reduced percentage of PKO. In order to explain the observed deactivation profile of Lipozyme IM TM during the continuous experiment, the effect of (i) initial water activity ( a ) of the biocatalyst and (ii) solvent washing of the lipase preparation on the catalytic activity were further investigated in batch transesterification reactions. 2. MATERIALS AND METHODS 2.1. Materials Refined, bleached and deodorized palm kernel oil (PKO) and palm oil stearin (POS), the obtained fraction of palm oil rich in satm'ated fatty acids, were supplied by FIMA, Produtos Alimentares, Portugal. The commercial preparation of the lipase from Rhizomucor miehei, "Lipozyme IM TM'', immobilized by adsorption on an anion exchange macroporous resin (0.2 to 0.6mm diameter), was kindly donated by NOVO Industries, Denmark. This is a thermostable 1,3- specific lipase preparation with a temperature optimum around 600C-70~ 2.2. Methods 2.2.1.Batch Transesterification Reaction Batch preliminary studies were carried out to optimize the blend initial composition, reaction temperature and biocatalyst load. The immobilized lipase was added to the reaction medium (70g) composed by different proportions of POS and PKO. The reactions were carried out inside small magnetically stirred thermostated cylindrical glass reactors (100 ml) at the desired temperature (higher than 45~ to prevent medium solidification). At different reaction times, 3 ml samples were taken out, paper filtered in an oven at approximately 80~ to remove biocatalyst particles, and the SFC was determined (cf. 2.2.6). 2.2.2. Optimization of Batch Transesterification Conditions
Temperature and Medium Composition The best reaction conditions for Batch Transesterification Reaction were established via the Response Surface Methodology (RSM) 3 as a function of both temperature and medium composition (concentration of POS).
437
A central composite rotatable design was used, with 6 replicates of the central point (14 experiments). POS concentration varied between 55% and 100%; temperature between 660C and 77~ and the biocatalyst load was fixed at 5% (w/w). Samples were taken at the following reaction times: 1, 2, 4 and 6 h. Results were analysed by the program "Statistica TM'', version 5, from Statsoft, USA. Tridimensional response surfaces, corresponding to the SFC3s.c values, were obtained for each of the considered reaction times. - Biocatalyst Load The effect of Lipozyme IM TM concentration on the reduction of SFC35.c was evaluated in order to gather more insight for the operation of the continuous reactor. The protocol was similar to the previously described (cf. 2.2.1.): POS initial concentration was 70% (w/w) and the temperature was 70~ 2.2.3. Continuous Transesterification Reaction The continuous transesterification reaction was carried out, for 5 days, in a continuous stirred tank reactor (CSTR), at 700C and a residence time of 15 minutes. The reactor was a magnetically stirred thermostated cylindrical glass vessel (100 ml) closed with a rubber stopper pierced by inlet and outlet glass tubes. The untransesterified fat (70% POS and 30% PKO) was continuously pumped at a flow rate of 5.2 ml/min, from a reservoir at 70~ through a silicone tubing to the bioreactor. Across the outlet glass tube, a sieve-like plate, made of porous glass Go (maximum pore size between 160 and 250 }.tm) prevented the biocatalyst to wash out. A load of 20% (w/w) ofLipozyme IM TM was used. 2.2.4. Treatment of the biocatalyst used in the continuous experiment After a 5 day continuous run, the enzyme preparation was removed from the bioreactor and divided into two portions: one of them was washed with n-hexane and the aw adjusted to 0.22 by contacting with the vapour phase of a saturated solution of potassium acetate, at 23~ for 4 days 4. The "treated" biocatalyst and the original one were reused in batch transesterification experiments, under the same reaction conditions as for the continuous experiment. 2.2.5. Reactions under an initially controlled water activity The influence of initial water activity (a~) of the biocatalyst on transesterification kinetics was investigated, since high a~ values promote the competing hydrolytic reaction 4's'6. The biocatalyst was previously (i) dried under vacuum at 40~ for 5min (final a~= 0.31), 15min (final a~= 0.20) or 30min (final a~=0.12) or (ii) equilibrated, for 4 days at 220C, with the vapour phase of saturated salt solutions of known aw- KCH3COO , (a~ = 0.23); K2CO3, (aw =
0.43); Mg(NO3) 2, (a~ = 0.53); NaC1, (aw =0.75). The final aw was assayed in a ROTRONIC HYGROSKOP DT humidity sensor (DMS100H). The immobilized lipase at an initial pre-established a~ value was used in batch transesterification reactions (cf. 2.2.1.).
438
2.2.6. Analytical methods Solid Fat Content Assay: The time course of the reactions was indirectly followed by measuring the Solid Fat Content (SFC) of the samples in a pulsed Nuclear Magnetic Ressonance (NMR) Spectrophotometer (Minispec P-20i, IBM). Each sample replicate was melted at 60~ for 5 minutes, followed by 60 minutes at 0*C and, at last, 30 minutes at 35~ Free Fatty Acids (FFA): FFA present in reaction media, due both to lipase-catalysed interesterifieation kinetics4 and hydrolysis of glycerides occurring even under low a~6'7'8, were assayed by titration with a 0.1N sodium hydroxide aqueous solution. The percentage (w/w) of FFA was calculated on the basis of the molecular weight of oleic acid.
3. RESULTS AND DISCUSSION The best reaction conditions for batch transesterifieation were established via the Response Surface Methodology (RSM). Transesterification reaction was significantly affected by medium composition (linear and quadratic terms). Temperature was not a significant effect in the range tested (66-77~ during the first 6 hours. This is in agreement with previously reported results 9. Further statistical analysis was carried out only on the significant effects. The tridimensional surface fitted to the experimental data points of 8FC35. c after 1 hour reaction time, was described by the following polynomial equation: SFC3s.c = 9.74 - 0.315 POS + 0.005 POS 2
(1)
The high values of both the determination coefficient (R2) and the adjusted determination coefficient (R.2,dj), 0.9751 and 0.9706, respectively, indicate the goodness of fit of this model. After 1 hour reaction-time, the observed decrease in SFC35.c values was similar or even higher than the results obtained by chemical transesterification. Best results (corresponding to final SFC35.c values lower than 12) were observed with 55 to 70% of POS. Similar profiles were obtained for results after 2, 4 and 6 hours reaction-time. This seems rather promising from an industrial point of view. A concentration of 15% (w/w) Lipozyme IMTM was selected for the continuous experiment since an important decrease in SFC3s,c values was observed after 15 minutes of batch transesterification time (Fig. 1). This is a suitable residence time for the continuous operation. Higher biocatalyst loads did not lead to further SFC35.c decrease during the same period. During the continuous experiment, a fast decrease in enzyme activity was observed (Table 1.), accompanied by an increase in the water activity (a~) of the biocatalyst (from 0.077 to 0.363). This fast deactivation of Lipozyme IMTM is in contrast with other experiments where the same biocatalyst was used either in solvent-free media ~~or in the presence of an organic solvent I~.
439
Fig. 1- The effect of biocatalyst 1o%[ concentration on the ---o-- 15%11 reduction of SFC3ro --6-- ~/,I during batch ...0_25%I transesterificationof 70% POS with 30% 35*/0] PKO, at 70~
2O r162 15 O
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A recovery of the catalytic activity was observed (Table 2.) when this enzyme preparation, washed with n-hexane and the a~ adjusted to 0.22, was reused in batch transesterification. This suggests a reversible inactivation probably ascribed to the presence of oxidation products in the microenvironment and/or variations in a~.
Table 1- SFCjrc values of the samples
taken during the continuous transesterification experiment Tim e (h) ,, SFC~5~(%) 0 20.32 0.75 17.40 4.5 18.00 16.5 17.86 28.5 19.36 40.5 19.06 52.5 20,45 64.5 20.36 76.5 19.61
Table 2- Performance of the biocatalyst previously
used in the CSTR, directly used in batch transesterification experiment ("Untreated biocatalyst") or after solvent wash and aw equilibration ("Treated biocatalvst"~ SFC3rc (%) SFC35*c (%) Reaction ("Untreated" ("Treated" Time (min) biocatalyst) biocatal),st) 0 24,20 22,24 15 22,78 14,51 30 21,94 12,81 45 22,01 10,51 . 60 21,83 9,50
Further studies were carried out to investigate the effect of the initial a~ of the biocatalyst on transesterification activity (Fig. 2). The methodology used for a~ adjustment (vacuum drying or pre-equilibrium with the vapour phase of saturated salt solutions) does not seem to affect the biocatalyst activity (Fig. 2). A rise in a~ was responsible for a decrease in the SFC3rc, which is in contrast with the results from the CSTR experiment.
440 !00 15 min
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Figura 2: SFC3rc values and FFA release during batch transesterification of
70%POS, at 70~ catalysed by "Lipozyme IM rM'' (20%, w/w) at different a~, attained by vacuum drying (n, o) or by contacting the enzyme preparation with the vapour phase of saturated salt solutions (l, o). In addition, an increase in FFA with the water activity was observed (Fig. 4). Activity recovery was probably due to the removal, by the solvent, of inhibitory compounds from the microenvironment, rather than a water activity effect. The FFA accumulation (4-18%, w/w) seems to be a limiting factor for scaling up this system. Acknowledgements- One of us (V. Falasehi) was partialy sponsored by the E.C. Socrates (Erasmus) Programme. REFERENCES
1- Erickson, M..D.E. (1995), In: Practical Handbook of Soybean Processing and Utilization, (D. R. Erickson, ed.), AOCS Press and United Soybean Board, pp. 277-296. 2- Faur, L. (1996), In: Oils and Fats Manual (A. Karleskind and J.P. Wolff, eds.), Assoc. Fran~.]~tude des Corps Gras, London & Paris, pp.923-925. 3- Montgomery, (1991)Design and Analysis of Experiments, John Wiley&Sons, 521551. 4- Ferreira-Dias, S.; Fonseca, M.M.R.(1995), Bioprocess Eng., 12 (5) 327-337. 5- Macrae, A.R. (1985), In: Biocatalysis in Organic Synthesis (J. Tramper, H.C. van der Plas, P. Linko, eds.), Elsevier, Amsterdam, pp. 195-208.. 6- Kyotani, S.; Fukuda, H.; Nojima, Y.; Yamane,T. (1988), J.Ferm.Technol. 66 (5) 567-575. 7- Heisler, A.; Rabiller, C.; Hublin, L. (1991), Biotechnol. Lett. (13) 327-332. 8- Osterberg, E.; Blomstrom, A.C.; Holmberg, K. (1988), J.Am.OilChem.Soc., 66 (9) 1330-1333. 9- Bloomer, S., Adlercreutz, P., Mattiasson, B. (1991), Biocatalysis, 5:145-162. 10- Forssr P., Parovuori, P., Linko, P., Poutanen, K. (1993), J.Am. Oil Chem Soc, 70 (11) 1105-1109. 11- Bloomer, S., Adlercreutz, P., Mattiasson, B. (1992), J.Am.Oil Chem Soc., 69 (10)
966-973.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
441
Effects of Lipid-Borne Compounds on the Activity and Stability of Lipases in Microaqueous Systems for the Lipase-Catalyzed Interesterification X. Xu a, C.-E. Hay b and J. Adler-Nissen a aDepartment of Biotechnology, Building 221; and bDepartrnent of Biochemistry & Nutrition, Building 224; Technical University of Denmark, DK-2800 Lyngby, Denmark In this paper, the effects of lipid-borne compounds such as lipid hydroperoxides, lipid polymers, phospholipids and emulsifiers, cholorophyll and carotenoids, tocopherols, citric acid, partial glycerides and free fatty acids on the activity and stability of lipases are discussed. Generally, the content of above compounds in commercial edible oil level did not affect the initial activity except for partial glycerides and free fatty acids, but the stability was influenced by most of the compounds. Besides tocopherols and partial glycerides, all the other minor compounds in oils and fats discussed had a tendency to reduce lipase life-time.
1. INTRODUCTION Lipases, so far, have not been well developed and applied in or by industry compared to some of the other enzymes, such as proteases, amylases, cellulases, etc., even though detergent application of lipases is growing very fast due to the developments of genetic engineering [1]. The whole enzyme business is still focusing on protein hydrolysis, starch modification, animal feed, baking industry, cotton fiber and textile processing, detergent and personal care [2]. The applications of lipases in fats and oils industry remain far behind. The partial reason is due to the need to find the competitive or value-added products. The other reason may be the cost or consumption of lipases and the efficiency of the process. If the applications are wanted for the normal lipid production or modification, the cost of the lipases has to come down and the stability of the lipases needs to be improved. Lipase research, whether fundamental or technological, on the other hand, arose great interest in the past decade [3]. One of the big advances has been the structural elucidation of lipases since 1990 [4]. This progress will soon give the detailed structural explanation of deactivation and inhibition of lipases and solutions to the problems will also be derived thereatter. Another major advance was the development of enzymatic catalysis in microaqueous media initiated by Klibanov and coworkers in 1984 [5]. This progress provides more possibilities of potential products for industrial application. It is convincing now that enzymes, if optimally applied, work as well in micro-aqueous media as in aqueous system [6]. Enzyme stability and activity inhibition were a long-time-concerned topic for the industrial applications. Lipase inhibitors in aqueous media were investigated in the past decades [7-8]. The effect of pH value and colipase on lipases were also investigated in both aqueous and non-aqueous media [9-10]. As the non-aqueous enzymology has attracted more
442 concems recently, the effects on the stability and activity of the enzymes have received some research, of solvent [11 ], water content [12], additives [13], as well as structural elucidation of thermal deactivation on the base of the protein engineering progress [14]. The broad understanding of enzymes and functional or property improvement can be expected in the near future. Lipid modifications are still the important and promising applications for lipases regardless of the slow progress compared to the detergent uses [1,9]. These applications include the modification of fats and oils and production of partial glycerides by interesterification and the enrichment of n-3 fatty acids from fish oil [1], etc. They are normally carried out in non-aqueous systems with or without solvent, though solvent-free systems are preferred in most of the applications if they are not absolutely necessary. The possible influences of impure compounds in the oils and fats had arisen concerns [ 15-17] and the effects of lipid peroxides [18-19] and phospholipids [20] on the stability and activity of lipases were also reported. In this report, all important minor compounds existing in the oils and fats are discussed. 2. LIPID HYDROPEROXIDES
Lipid hydropcroxides are formed from lipidauto-oxidation and photo-oxidation, which often occurred in industrialoils,only differingin theirextent [21]. Lipid hydroperoxides were reported to bc toxic to the human body because of their possible inhibition to the enzymes [22]. The content of lipid hydroperoxides is empirically measured as peroxide value (POV), which is reported in units of milli-equivalentsof oxygen per kilogram of oil. The inhibition and inactivationof lipasc,in vitro,by fat peroxides were firstreported by Ohta et al. [18] in the course of batch and continuous glycerolyses of fat catalyzed by free Pseudomonas fluorescens lipase.The initialactivityof the lipase linearlydecreased in the batch system. The stabilityof the lipase in the continuous membrane reactor was also greatly decreased by the increase of P O V of the oil.The reason of deactivation, from the same article,was caused by the polymerization of the lipase due to the existence of hydroperoxides. In another paper, W a n g and the coworker [19] used immobilized Lipozyme IM (Mucor miehei) for the interesterificationbetween soybean oil and laurie acid in the batch reactor with petroleum ether as the solvent. They found that the initialrate was not affectedby the differentvalues of the oil,but stabilityof the lipase was reduced. Thus they suggested that P O V of the oils more than 5 meq/kg would strongly reduce the useful lifeof the enzymes. In the reaction between tcaseed oil and the mixture of stearicand palmitic acids in hexanc catalyzed by immobilized porcine pancreatic lipase,the conversion of the reaction using a tcaseed oil (POV 27meq/kg) was only about a half of that of the oil in normal edible use (POV 3 meq/kg) after 4 batches (12 hours each batch) [23]. In the production of structured lipids between sunflower oil and capric acid in a pilot enzyme bed reactor using Lipozyme IM as the biocatalyst,the effectof lipid oxidation products on the stabilitywas also observed in a system without solvent (Fig. l). The hydroperoxide compounds were found almost to be fully absorbed by the enzyme bed because the effluents from the bed had a P O V of zero [23]. This phenomenon will, of course, probably be affected by other process parameters and the carder property of immobilized enzymes.
443 3. PHOSPHOLIPIDS AND EMULSIFIERS
Phospholipids are one of the very important minor components in vegetable oils. The content of phospholipids in crude vegetable oils normally varies from 0.4 to 3.5% [21]. After refining, the content is normally less than 40 ppm. Sometimes phospholipids are also added in the oils as antioxidants in ppm level [22]. Emulsifiers, sometimes also called detergents or surfaetants, are added to oils for different purposes. Normally shortening oils contain a little emulsifier [21] and some amphiphilic substrates are added in organic system to form reversed micelles.
100 A
[
100
.
.
.
.
.
90
90
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50
50 40
Running time (days)
Fig. 1. Effect of lipid hydroperoxides on the stability of Lipozyme IM. Experiment conditions: enzyme bed 87 (1) x 5 (i.d.) cm; substrate ratio, eapric acid/sunflower oil (6/1 mol/mol); temp., 60~ residence time, 146 min; water content, 0.12%. POV value of sunflower oil as 4.2 (*) and as 24.7 meq/kg ( ) .
,
Running bme (days)
Fig. 2. Effect of lipid polymers on the stability of Lipozyme IM. Experiment conditions: enzyme bed 91 (1) x 5 (i.d.) em; substrate ratio, capric acid/sunflower oil (2/1 mol/mol); temp., 60~ residence time, 95 min; substrates saturated with water. Polymers content of sunflower oil as 0.2 (*) and as 12.4% ( ) .
There were seemingly controversial reports regarding the effects or functions of above compounds in the lipase-catalyzexi reaction system. Some reports used free or immobilized tracylglycerol lipases to hydrolyze or modify the phospholipids and very good yields were claimed [24-25]. The uses of detergents and bis(2-ethylhexyl) sodium sulfosuccinate (AOT) to form reversed mieelles were also reported to increase the initial activity of lipases in the organic solvent system [26-27]. On the other hand, detergents were reported to be inhibitors [7]. The inhibition of AOT was also reported [28]. The effect of phospholipids on the activity and stability of Lipozyme IM in the organic media and batch system was found very crucial [20]. The authors suggested that the oils used for reaction should have less than 200 ppm phospholipids if unacceptable inactivation of Lipozyme IM was to be avoided. From the structural point of view, lipases are inhibited by most amphiphilic compounds because of inteffacial adsorption [29]. In our working system, we found that phospholipids were totally absorbed by the enzyme bed during the first five days [23]. If this
444 happens continuously in the whole life of the enzyme, even 40 ppm phopholipids will eventually be a strong factor on the stability of the lipase. 4. PIGMENTS
Crude vegetable oils contain small amount cholorophyll (green) and carotenoids (yellow or red) depending on oil sources, oilseed quality and production methods [21 ]. After bleaching, green color normally does not exist. The fully refined oils are normally light yellow depending largely on refining processes. However, virgin olive oil, a traditional Mediterranean oil, is normally greenish and red palm oil, a new developed nutritional palm oil, is almost red. The effects of cholorophyll and carotenoids on the stability of lipases have not been reported. In one of the experiments, we found that both the pigments reduced the stability of the porcine pancreatic lipase which was immobilized on diatomaceous earth by physical absorption. Also the green color of degunmaed olive oil had mostly disappeared and the color of red palm oil was also lighter after the reactions [23]. This means that the pigments were fully or partially retained and absorbed by the immobilized enzyme.
5. LIPID POLYMERS
Lipid polymers exist in oils and fats after oxidation or thermal treatments such as frying or deodorization. Refined oils and fats normally contain less than 1% dimers, trimers and polar triglycerides [30]. Fried oils may contain up to 10% polymerized compounds [31 ]. A few publications reported the production of special polymers by enzyme-catalyzed reactions [33]. Polymers in the enzyme-catalyzed hydrolysis system were also reported to increase the activity in the special situation [32]. In our study, the effect of polymers on the stability of the lipase was observed (Fig. 2). The activity was gradually decreased after feeding the fried sunflower oil with 12% polymers and the POV of 9 meq/kg. The effluents contained only 1.3% polymers analyzed by size-exclusion chromatography [23]. Polymers can also be retained by the enzyme bed used, especially in the first 20 hours of running.
6. ANTIOXIDANTS Edible oils normally contain small amounts of antioxidants for preventing their autooxidation. Natural antioxidants normally include tocopherols, phospholipids, ascorbic acid (Vitamin C), phytic acid, phenolic acids and so on. Common synthetic antioxidants for edible use are butylated hydroxyanisole (BHA), butylated hydroxytoluene (BHT), propyl gallate (PG), tertiary butyl hydroquinone (TBHQ), etc. [22]. Tocopherols were reported to prevent the thermal inactivation [33]. However, citric acid and phytic acid were reported to be inhibitors to lipase activity [34]. In our study, two commercial antioxidant products were investigated for the effect on lipase stability [23]. Grindox Toco 50, a natural tocopherol concentrate from Danisco Ingredients, Denmark, did not affect the stability of the immobilized lipase in the first ten days by adding 400 ppm in the feeding substrates containing 50% refined fish oil and 50% capric acid in a enzyme bed reactor similar to those in figure 1 and 2. Grindox 117, a product
445 containing 17.5% PG, 10.0% citric acid, 7.5% aseorbyl palmitate and 65.0% mono-and diglycerides of fatty acids and propylene glycerol from the same company, on the other hand, was found to reduce the 18% interesterification activity compared to the control after 10 days by adding 200 ppm in the substrates in the same system. It is difficult to say, however, which compound plays the main role. The elucidation of main antioxidants on the stability is under way.
7. DISCUSSION There are more minor compounds in oils and fats than described above, depending on the oil sources [21 ]. For example, heavy metal ions existing in oils and fats in ppb level will certainly effect the stability of enzymes [17]. Different free fatty acids had different incorporation in the micro-aqueous interesterification system [35]. The inhibition of dodecanoic acid in the olive oil hydrolysis was also reported [36]. As can be seen from above discussions, enzyme bed reactor which was promising for lipase-catalyzed interesterification [37], acted as a chromatography column to retain the more polar or complex compounds. In batch stirred tank reactors, polar compounds will also be absorbed on the enzyme surface. The inhibition happens, probably, because all these minor compounds covers the surface of the enzyme and eventually prevent the contact between the enzyme and the substrates, except lipid hydroperoxides, or possibly, heavy metal ions, which react with enzymes and cause the conformational changes. Certainly, the retaining of the minor compounds by the enzyme bed will differ with different hydrophobicity of the substrates. When shorter chain length free fatty acids are used as acyl donors, the less retaining of the polar compounds can be expected. Preliminary experiments were carded out to improve the enzyme activity after partial inhibition by lipid hydroperoxides, lipid polymers (fried oil), cholorophyll, carotenoids and antioxidants (Grindox 117) by feeding the mixture of hexane, diethyl ether and ethanol (70/20/10 VN/V) to clean the lipase. Roughly 20%, 40%, 37%, 87% and 70% of the lipase activities were recovered compared to those of the controlled experiments, respectively, of the inhibition caused by above-mentioned compounds [23]. It seems that lipid hydroperoxides, cholorophyll and lipid polymers (flied oil) have stronger interactions with lipases and the inhibitions are mainly irreversible. The activities of the others, more or less, can be recovered and the inhibition is mainly caused by covering the lipase surface and limiting the mass transfer from the bulk solution to the lipase active sites of the substrates. The partial loss of activity may also be due to the inhibition of the solvent used [ 11 ]. Generally, the impurities in oils and fats, whether more or less, will eventually cause the reduction of the lipase life, especially in the enzyme bed reactor and micro-aqueous solvent-free systems. The quality of oils and fats have to be improved, as much as possible because lipases now are still relatively expensive. In the short run, the cost of lipases will come down with the development of genetic engineering. The cost of oil refining and enzyme consumption will reach a balance. A compromise has to be made regarding the oil quality and lipase consumption for optimal economies of the processes.
Acknowledgment The financial support from the Danish Technical Research Council is appreciated (project No.: 9601280). Partial work was previously done by XX in Zhengzhou Grain
446 College, China. We thank T.T. Hansen and M. W. Christensen, Novo Nordisk A/S, for supplying lipases and other helps. Hong Zhang is thanked for doing polymer analysis. References 1. P. Eigtved, Enzymes and lipid modification in: Advances in Applied Lipid Research (ed. by F. B. Padley), vol. 1, JAI Press Ltd, London, 1992. 2. Annual Enzyme Business Report (1997) of Novo Nordisk A/S, Denmark (Available at the web site: www.novo.dk). 3. F.X. Malcata (ed.), Engineering of/with Lipases, Kluwer Academic Publishers, Dordrecht, 1996. 4. R.L. Brady, A.M. Brzozowski, et al., Nature, 343 (1990) 757-770. 5. A. Zaks and A.M. Klibanov, Science, 224 (1984) 1249-1251. 6. A.M. Klibanov, Trends in Biotechnol., 15 (1997) 97-101. 7. H. Zollner, Handbook of Enzyme Inhibitors, VCH, 1989, p211. 8. S. Patkar and F. Bjorlding, Lipase inhibitors, in: Lipase: Their Structure, Biochemistry and Application, Cambridge University Press, 1994. 9. W.J.J. van den Tweel, A. Harder, et al. (eds.), Stability and Stabilization of Enzymes, Elsevier Science Publishers B. V., 1993, pp13-20, 111-131,445-450. 10. L. (3. Copping, R.E. Martin, et al. (eds.), Opportunities in Biotransformations, Elsevier Applied Science, 1990, pp81-87. 11. P. Aldercreutz and B. Mattiasson, Biocatalysis, 1 (1987) 99-108. 12. G..Bell, P. Hailing, et al., Trends in Biotechnol., 13 (1995) 468-473. 13. A. O. Triantafyllou, E. Wehtje, et al., Biotech. Bioeng., 54 (1997) 67-76. 14. S. E. Zale and A.M. Klibanov, Biochemistry, 25 (1986) 5432-5444. 15. L.H. Posorske, G.K. LeFebvre, et al., J. Am. Oil Chem. Soc., 65 (1988) 922-926 16. R. A. Wisdom, P. Dunnill, et al., Biotechnol. Bioeng., 29 (1987) 1081-1087. 17. C. Ratledge and S.G. Wilkison (eds.), Microbial Lipids (2), Academic Press, 1989, pp637-641. 18. Y. Ohta, T. Yamane, et al., Agric. Biol. Chem., 53 (1989) 1885-1890. 19. Y. Wang and M.H. Gorden, J. Agric. Food Chem., 39 (1991) 1693-1695. 20. Y. Wang and M.H. Gorden, J. Am. Oil Chem. Soc.,68 (1991) 588-590. 21. X. Xu, T. Yang, et al., Oil and Fat Chemistry, China Commercial Publishing House, 1993. 22. D. L. Madhavi, S.S. Deshpande, et al. (eds.), Food Antioxidants, Marcel Dekker, Inc., 1996, pp38-39, 41-52 and 67-86. 23. X. Xu, unpublished results. 24. M. Ghosh and D.K. Bhattacharyya, J. Am. Oil Chem. Soc., 74 (1997) 597-599. 25. I. Svensson, P. Aldercreutz, et al., J. Am. Oil Chem. Soc., 69 (1992) 986-991. 26. {3. M. DeUamora-Ortiz, R.C. Martins, et al., Biotechnol. Appl. Biochem., 26 (1997) 31-37. 27. D. M. F. Prazeres, et al., Biotech. Bioeng., 41 (1993) 761-770. 28. C. Otero and L. Robledo, Progr. Colloid Sci., 98 (1995) 219-223. 29. D. W. S. Wong, Food Enzymes, Chapman & Hall, 1997, pp170-211. 30. F. D. Gunstone, J.L. Harwood, et al. (eds.), Lipid Handbook (2~ ed.), Chapman & Hall, London, 1994, pp249-318. 31. G. M~rquez-Ruiz, G. Guevel, et al., J. Am. Oil Chem. Soc., 75 (1998) 119-126. 32. R. Arroyo, F. J. S~nchez-Muniz, et al., J. Am. Oil Chem. Soc., 74 (1997) 1509-1516. 33. J. S. Dordick (ed.), Biocatalysts for Industry, Plenum Press, New York, 1991,pp83-112, 241-255, 161-180. 34. E. T. Champagne and R.J. Hron, Cereal Chemistry, 71 (1994) 483-487. 35. J. Tramper, H.C. van der Plas, et al. (eds.), Biocatalysts in Organic Syntheses, Elsevier Science Publishers B.V., Amsterdam, 1985, pp 195-208. 36. M. Markweg-Hanke, S. Lang, et al., Enzyme Microb. Technol., 17 (1995) 512-516. 37. X. Xu, S. Balchen, et al., J. Am. Oil Chem. Soc., 75 (1998) in press.
Stability and Stabilizationof Biocatalysts A. Ballesteros,F.J. Plou, J.L. Iborra and P.J. Halling (Editors) 9 1998Elsevier Science B.V. All rights reserved.
447
Stabilisation of lipases for activity in ammoniolysis F. van Rantwijk, A.C. Kock-van Dalen and R.A. Sheldon* Laboratory of Organic Chemistry and Catalysis, Delft University of Technology, Julianalaan 136, 2628 BL Delft, The Netherlands 1.
INTRODUCTION
Enzymes that are used in organic media are usually immobilised for efficiency. Moreover, adsorption or multi-point attachment of enzymes may increase their stability under low-water conditions. The lipophilic nature of lipases favours their adsorption on non-polar surfaces, a feature that has been exploited in their immobilisation on macroporous polypropylene (Accurel | EP100) for use in transesterification [1,2,3,4]. It has also been shown that adsorption on EP100 increases the catalytic efficiency of P. cepacia lipase for hydrolysis in organic media [5]. Porous polyurethane foams that are formed in situ from prepolymers have the potential advantage that the enzyme is stabilised by multi-point attachment. The method has been used for chymotrypsin [6], ~-galactosidase [7] and phosphortriesterase [8], but not for lipases. We have investigated the stabilisation of lipases for the ammoniolysis of carboxylic acid esters to amides [9] by adsoption on polypropylene as well as by entrapment in polyurethane foam and now present a preliminary report. 2.
EXPERIMENTAL
2.1. M a t e r i a l s a n d m e t h o d s Lipases from Candida antarctica (B type, SP525) and Humicola sp. (SP 523) (kindly donated by Novo Nordisk A/S) were used as received. Pseudomonas alcaligenes lipase (Lipomax) was received from Gist-brocades B.V. as a gift; it was suspended in distilled water containing 5 mM calcium gluconate and centrifuged; the supernatant was assayed for lipase activity and used in immobilisation experiments. Pseudomona mendochina lipase (Lumafast 2000) was donated by Genencor International and was subjected to the same treatment before use. Batches of Accurel EP100 (0.4 - 1 mm) were kindly donated by Akzo Nobel Faser A.G. HYPOL 3000 and 5000 prepolymers were gifts from Hampshire Chemical Ltd. and were used as received. Full experimental details will be reported elsewhere. Soluble as well as immobilised lipases were assayed in a standard tributyrin
*
Author to whom correspondence should be addressed.
448 hydrolysis test. The progress of the reaction was monitored by automatic titration. 1 Unit (1 U) liberates 1/~mol of butyric acid per minute. Ammoniolyses of ethyl dodecanoate were carried out as described [9] and were monitored by GC. 2.2. I m m o b i l i s a t i o n Accurel EPIO0. The carrier material was treated as described [3] and shaken for 16 h with a solution of lipase. A sample of the supernatant was taken and assayed for lipase activity. The immobilised enzyme was filtered off, rinsed with water and dried in vacuo for 16 h at 20 ~ In the titration experiments the carrier was added in portions, a sample of the supernatant was taken after a suitable period and assayed for lipase activity. Polyurethane foam entrapment of the lipases was carried out as described [8] using HYPOL prepolymer in HEPES buffer containing Brij 52 and CoC12.
3.
R E S U L T S AND D I S C U S S I O N
3.1. A d s o r p t i o n o n A c c u r e l E P 1 0 0 Initial adsorption experiments with C. antarctica lipase revealed that the amount that could be adsorbed on Accurel EP100 was low compared with literature data [3] and previous results obtained at our lab [10]. Scanning electron microscopy (SEM) of the carrier material showed that its cell size had increased by a factor of 4 5 between 1992 and 1995, with a corresponding reduction of the surface area.
Figure 1. SEM photographs of two batches of Accurel EP100 at 400 x magnification; A: batch 7/92, B: batch 7/95. Solutions of the lipases from C. antarctica, P. alcaligenes and P. mendochina were titrated with Accurel EP100 (new batch). The loading limit of C. antarctica lipase was 27 kLU per g EP100 and P. mendochina lipase was adsorbed to a limit of approx 23 kLU/g. Titration of P. alcaligenes lipase with EP100 resulted in an unexpected S curve when the loading of the carrier was plotted against the lipase concentration in solution (Fig. 2). The reason for this effect is obscure but formation
449 of enzyme aggregates at > 8 kLU/ml would seem a likely explanation. 800 c~ 600 ...I
c~ 400 r -o
o
...I
200 b',,,,
| ,,,
, l,
0
,,,,,
, ,,
2
i,,
,,,,,,'l
i,
4
6
Concentration
i,
,,i,
i,i
,i
8
,,,
l,,,
I
10
(kLU/ml)
Figure 2. Loading of P. alcaligenes lipase on EP100 as a function of concentration Table 1. Activity and leaching tests of immobilised lipases Preparation a
Loading (kLU/g)
C. antarctica
26
Novozym 435 c
unknown
Found (kLU/g) 3.1
Yield (%) 12
4.1
Activity in filtrate (kLU/g) b 0.3 1.8
P. alcaligenes
18
2.0
11
n.d.
P. alcaligenes
37
3.3
9
n.d.
P. alcaligenes
190
4.7
2.5
0.9
196
2.9
1.5
n.d.
P. alcaligenes
d
P. mendochina
e
15
0.9
6.1
n.d.
P. mendochina
f
19
0.5
2.7
n.d.
a. On EP100 unless noted otherwise; b. per gram of preparation used; c. on a proprietary carrier; d. carrier pretreated with albumin [11]; e sorbitol (0.5 M) added; f. sorbitol (0.1 M) added. In order to check the efficiency of the adsorption method, the immobilised lipases were dried and subsequently assayed in tributyrin hydrolysis tests (Table 1); only a fraction of the adsorbed lipase was still active, as has been reported previously [2]. P. mendochina lipase required the addition of sorbitol to the immobilisation buffer to maintain its activity after drying. Hydrolysis tests of adsorbed lipases are indicative only, since the reaction continued after filtration at up to 40 % of the previous rate due to redissolved lipase.
450 Table 2. Ammoniolysis of ethyl dodecanoate Enzyme preparation
a
Loading
Amount
(kLU/g)
(g)
Ammoniolysis (h/% reaction) 1
2
5
24
48
,
C. antarctica/EP 100
27.5
Novozym 435
unknown
P. alcaligenes
P. mendochina/EPlO0
0.04
18
28
50
85
0.04
21
33
57
93
20
30
45
28
46
76
1400
0.01
190
0.04
15
0.10
P. alcaligenes/EPlO0 b
,.
19
29
43
a. Reaction conditions: 2 mmol ethyl dodecanoate in 4 ml t-butyl alcohol, 2.5 M NH3 at 40 ~ b. immobilised in the presence of 0.5 M sorbitol. Some representative samples of the immobilised lipases were assayed in the ammoniolysis of ethyl dodecanoate (Table 2). Of these, C. antarctica lipase was by far the most active one on a per-unit basis. The example of P. alcaligenes lipase clearly shows the effect of adsorption on the catalytic efficiency in an organic medium: freely suspended enzyme was 7 times less active - on a per-unit basis - t h a n the one adsorbed on Accurel EP100. The P. mendochina enzyme was less active t h a n the one from P. alcaligenes, but it suffers mainly from the practical disadvantage that only low a m o u n t s are adsorbed on Accurel EP100. Table 3. Ammoniolytic activity of P. alcaligenes lipase on Accurel EP100 Loading
Carrier
Amount a
(kLU/g)
pretreatment
(kLU)
330
-
190 196
Ammoniolysis (h/% reaction) 1
2
5
24
48
13.2
33
44
60
88
95
-
7.6
19
28
46
76
n.d.
albumin
7.8
23
35
56
87
94
37
-
1.5
7
10
16
40
60
35
albumin
1.4
6
11
22
51
75
a. Based on loading. We have briefly investigated the effect of some immobilisation parameters on the
451 ammoniolytic activity of P. alcaligenes lipase (Table 3). A low loading of the carrier resulted in some increase in specific activity, which was further increased by pretreatment of the carrier with albumin [11]. In conclusion the adsorption on Accurel EP100 increases the catalytic effmience of lipases in ammoniolysis but the rather low activity of the adsorbed lipases in hydrolysis tests indicates that there is much room for improvement.
3.2. Inclusion in p o l y u r e t h a n e foam Entrapment ofP. alcaligenes lipase in polyurethane foam was essentially carried out according to the published method [8]. We mainly used the Hypol 3000 prepolymer because it yielded a more flexible and hydrophilic foam than Hypol 5000, although the latter material gave a better efficiency (vide infra). Because only a few % of the activity could be recovered from the wash liquid (data not shown) it would seem that the remainder is tightly bound to the polymer. The hydrolytic activity of the resulting preparation was very low, however, corresponding to a few % of immobilisation yield. Attempts to increase the immobilisation effmiency by protecting the active site with a competitive inhibitor (p-nitrophenyl butylcarbamate) did not improve the yield. In ammoniolysis the foam-entrapped enzyme performed much worse than adsorbed enzyme (Table 4). Table 4. Inclusion of P. alcaligenes lipase in polyurethane foam Amount used (kLU)
Theor. loading
Activity
Yield
(kLU/g)
(U/g)
(%)
Ammoniolysis (h/% reaction) a kLU used
24
96
26 b
6.2
75
1.2
150 b
25.5
570
2.2
28
70
91
150 c
19.4
74
0.4
20
29
71
76
n.d.
P. alcaligenes lipase on EP100 (190 kLU/g)
7.6
a. Reaction conditions see Table 2.; b. HYPOL 3000; c. HYPOL 5000. Rather similar results were obtained with Novo SP523 (Humicola sp.) lipase (Table 5). With this enzyme a drop in immobilisation yield was observed when the loading was increased to a realistic level. A similar concentration dependence has been reported for a ~-galactosidase [7] but at a higher overall effmiency. The ammoniolytic activity was lower by a factor of 4 - 5 compared with a similar amount of enzyme adsorbed on EP100. In conclusion entrapment of lipases in polyurethane foam seems to result in their deactivation, contrary to results obtained with other types of enzymes by others [6,7,8] as well as by us [12]. It seems likely that deactivation is caused by carbamoylation of the active serine by the isocyanate function in the prepolymer.
452 Table 5. Inclusion of Humicola lipase in polyurethane foam Amount used (kLU)
Theor. loading
Activity
Yield
(kLU/g)
(U/g)
(%)
a
Ammoniolysis (h/% reaction) b kLU used
48
144
1
0.25
20
8
0.25
< 1
< 1
10
2.5
50
2
2.5
< 1
< 1
25
3
6
25
17
26
100
25
175
Humicola lipase on EP100 (300 kLU/g)
0.7
a. HYPOL 3000; b. reaction conditions see Table 2. 5.
ACKNOWLEDGEMENTS
Generous donations of enzymes by Novo Nordisk A/S (Bagsvaerd, Denmark), Gist-brocades (Delft, The Netherlands) and Genencor International (Delft, The Netherlands) are gratefully acknowledged. The authors wish to thank Akzo Nobel Faser AG (Obernburg, Germany) for gifts of Accurel EP100. HYPOL prepolymers were kindly donated by Hampshire Chemical Ltd. (Middlesbrough, UK) to whom the authors express their thanks. This work was supported by a grant from the Netherlands Ministry of Economic Affairs and carried out in cooperation with Chemferm Industrial Pharmaceuticals. REFERENCES
1. 2. 3. 4. 5. 6. 7.
A.R. Macrae, J.A. Bosley, A.D. Pellow, Eur. Pat. Appl. 322 213 A1. J.A. Bosley, A.D. Pellow, J. Am. Oil. Chem. Soc. 74 (1997) 107. S. Pedersen, P. Eigtved (Novo Nordisk A/S), PCT Int. Appl. WO 90/15868. B. AI-Duri, Y.P. Yong, J. Mol. Catal. B: Enzymatic 3 (1997) 177. G. Pencreac'h, J. Baratti, Appl. Microbiol. Bioteclmol. 47 (1997) 630. C.I. Mekras, M.H. George, J.A. Battle, Int. J. Biol. Macromol. II (1989) 2113. Z.-C. Hu, R.A. Korus, K.E. Stormo, Appl. Microbiol. Bioteclmol. 39 (1993) 289. 8. K.E. LeJeune, A.R. Russell, Bioteclmol. Bioeng. 51 (1996) 450; ILE. LeJeune, A.J. Mesiano, S.B. Bower, J.K. Grimsley J.R. Wild, A.R. Russell, Biotechnol. Bioeng. 54 (1997) 105. 9. M.C. de Zoete, A.C. Kock-van Dalen, F. van Rantwijk, R.A. Sheldon, J. Chem. Soc., Chem. Commun. (1993) 1831; M.C. de Zoete, A.C. Kock-van Dalen, F. van Rantwijk, R.A. Sheldon, Ann. N.Y. Acad. Sci. 799 (1996) 346. 10. M.C. de Zoete, unpublished results. 11. J.A. Bosley, Eur. Pat. Appl. EP 424 130 AI. 12. F. van Rantwijk, A.C. Kock-van Dalen, R.A. Sheldon, paper in preparation.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
453
W a t e r sorption i s o t h e r m as a tool to explore h y d r a t i o n of the m i c r o e n v i r o n m e n t of biocatalysts J. M. SS.nchez-Montero, R. M" de la Casa and J. V. Sinisterra Department of Organic & Pharmaceutical Chemistry. Faculty of Pharmacy. Universidad Complutense. 28040 Madrid. Spain The water sorption isotherm allows us to describe the hydration of the microenvironment of the biocatalysts in water immiscible organic solvents. The methodology is applied to lipases from C.antarctica and C. rugosa, purified, chemically modified or covalently immobilized on SiO 2. The influence that the nature of each solvent has and the biocatalyst in the shape of the isotherm is discussed. Using these isotherms, we can predict the a~ pre-equilibrium value of a reaction mixture to achieve the maximum yield in the esterification of unnatural substrates catalyzed by this biocatalyst in organic media. 1. INTRODUCTION The systematic study of enzymatic reactions in organic media is growing rapidly with the hypothesis of enzymes, as "on the shelf reagents " in organic chemistry [ 1]. In these systems, water plays a crucial role controlling the enzymatic activity, the thermal stability, the enantioselectivity and overall yield of product, particularly in reaction where water is produced such as esterification [2]. To carry out these processes on a big scale and/or long reaction time, the enzymes are stabilized by chemical modification or immobilized on supports. In these conditions the amount of water in the microenvironment of biocatalysts, considerably varies depending on the nature of the biocatalyst. This finding may simply reflect differences in the water distribution among all the components of the system: the organic solvent, the enzyme and the solid support [3]. Some authors have reported that immobilized enzymes on hydrophobic supports display better synthetic activity than those immobilized on hydrophilic carriers [4]. But lyophilization can dramatically reduce the synthetic activity of lipases immobilized on supports, such as agarose, silica and alumina. However, the activity of the lyophilized inorganic derivatives cannot be replaced when some water is added to the medium [5]. To understand the role of the water, it is necessary to develop simple and precise methodologies of analyzing the water distribution in the microenvironment of the biocatalysts and in the organic solvent. Recently, researchers have reported that reaction rates are a function of the water activity (a~) rather than of the water content of the medium. With immobilized lipases employed in an esterification reaction, Hailing et al. found similar shaped reaction rate-a~ profiles with most of the supports [6]. The water sorption (adsorption(solid)/absorption (liquid)) isotherm indicates the amount of water that can be adsorbed/absorbed, respectively, by a previously dehydrated sample as a
454 function of the water activity, a~. The methodology described in this paper is completely general and can be used with native lyophilized enzymes, and with immobilized (adsorbed or covalently bonded) or chemically modified enzymes. C.antarctica and C.rugosa lipases were used, purified, chemically modified or covalently immobilized on silica. 2. EXPERIMENTAL.
2.1. Enzymes and substrates Commercial lipase of Candida rugosa Type VII (CCRL) (8.7 IU/mg sample using olive oil as substrate) was from Sigma. Semipurification of Candida rugosa lipase was achieved by dialysis using a cut off membrane (20 kDa) as previously described [7]. CCRL was purified according to the method described by Rua et al. [8] and two isoenzymes were obtained: Lipase A (CRLA) and Lipase B (CRLB). Enzymatic activity using olive oil as the substrate: semipurified lipase (CRSL)=4.7 IU/mg solid); pure isoenymes, CRLA= 253 1U/mg protein; CRLB=290 IU/mg protein, Pure lipase B from Candida antarctica (SP525) was a gift from Novo Nordisk Bioindustrial S.A. Madrid. Racemic 2-(4-isobutylphenyl)propionic acid was kindly supplied by Boots Pharmaceuticals (Nottingham, U.K.). 1-propanol was obtained from Sigma Chemical. Isooctane (with analytical grade) and silica were from Merck. 2.2. Chemical modification The monomethoxypolyethyleneglycol (Mw = 5,000) (PEG) activated by pnitrophenylchloroformiate was obtained according to the procedure described by S~inchezMontero et al. [9]. The modification of the semipurified lipase from C. rugosa (CRSL) was performed at 4~ for 2 hours with gentle stirring in the standard buffer (0.1 M Tris/HCl buffer pH=8.0). The molar excess of activated PEG related to amino groups of the protein were tested (5/1 and 30/1) obtaining the chemically modified derivatives CRSL-PNFCF-1/5 and CRSL-PNFCF-1/30, respectively. The content of free lysines, before and after modification was determined using the procedure described by Snyder et al. [ 10]. The modification degree was CRSL-PNFCF-1/5 = 60% and CRSL-PNFCF-1/30 = 77% of the total lysines.
2.3. General Procedure for Immobilization. The activation of silica with 2,4,6-trichloro- 1,3,5-triazine (TCT) was carried out according to the methodology previously described by Moreno and Sinisterra, [ 11 ]. The immobilization of lipase was performed at 4~ 6 hours at slow stirring. One gram of silica was added to an enzyme solution (20 mg of CALB in 10 ml of Tris/HC10.1N (pH=8.0) buffer). After the desired contact time, the insoluble derivative was filtered and washed with standard buffer. Enzyme loading 15.6 mg of CALB/g of solid; 38% retained activity in the hydrolysis of tributyfin. 2.4. Measurement of adsorption isotherms. 200 mg of solid lipase (native or chemically modified) or immobilized derivative were first pre-dried with P2Os.The a~ values of solid preparations were measured at 25~ using a hygrometric sensor (Rotronic Hygroscopic D.T.) precalibrated with two saturated salt solutions at a~ = 0.11 and 0.98. The sorption isotherms in isooctane were measured with the same amount
455 of solid plus 1 ml of solvent. 2.5. General Procedure for Esterification. The reaction mixture was composed of isooctane (5 ml), racemic 2-(4isobutylphenyl)propionic acid (66 mM) and 1-propanol (66 mM). The reaction was started by adding different amounts of the lipase (native or chemically modified) to the solution and pre-equilibrating the system (solvent+biocatalyst) at a desired a~ value using a hygrometric sensor. The reactions were carried out at a fixed temperature by shaking 25 ml-flasks for a specified time. Then 100 pl of the solution was added to 1.4 ml of isooctane to analyze the ester conversion by gas chromatography. Candida rugosa lipase stereoselectively esterifies the (S) acid independently of the purification degree. Candida antarctica lipase stereoselectively esterifies the (R) acid.
2.6. Gas Chromatography Analysis. Gas chromatography analysis was performed in a Shimadzu GC-14A gas chromatograph equipped with FID detector, a split injector (1:2) and a SPB-1 sulfur column 15m• (Supelco, Bellafonte, PA, USA). Injector temperature was 300~ and the detector temperature 350~ carrier gas was nitrogen. Conditions for quantitative analysis were a column temperature of 180~ and a N 2 stream of 12 ml/min. 3. RESULTS AND DISCUSSION In Figure 1, we show the isotherms of CCRL in air and CCRL in isooctane (Fig 1A) and in MIBK (Fig 1B) they are compared to the water absorption isotherm of the only organic solvent. These solvents were selected as hydrophobic and hydrophilic solvent models. The isotherms of pure components cannot be used to describe the hydration level of the microenvironment of the solid biocatalyst that is different from the macroenvironment (the solvent)[2] because the water sorption of isotherm of this system (lipase + solvent), is different to the isotherms of the only solvent and of the dry lipase alone in the air. g water/g sample
g sample/ml solvent
9) CCRLin air 0.20 0.80 , ,.-- A. . CCRL+l:ooctam ' - ~ooctam t ~,,,i A I Isooctan 0.40 I
/~~
0.10
g water/g sample
1.20 -
e
0.80 B -
g sample/ml solvent -I CCRLin air
CCRL+MIBI
~
.I,K
1
~0.10
0.40 -
-I
.
0.00
r
0.00
-'
i
0.50 Aw
'
~0.00 1.00
0.00 1"0.00
0.20
"-
I 0.50 Aw
'
I o.oo 1.00
Figure 1. Water sorption isotherms of CCRL in air, of CCRL in the solvent and of the only solvent. Fig 1A: In isooctane. Fig 1B: In methyl isobutyl ketone. (MIBK) Thus, if we want to analyze the affect of the water on the microenvironment of the biocatalysts, the water sorption isotherm of the humected biocatalysts may be obtained.
456 The shape of the curves are related to the structure and physicochemical properties of the enzyme as we show in Figure 2 for CRLA (Figure 2A), CRLB (Figure 2B) and for lipase B of Candida antarctica CALB (Figure 2C). We can see that CRLA and CRLB show different sorption isotherms. The dry isoenzyme adsorbs more water the wet isoenzyme. This is explained by the presence of neutral sugars, 8% CRLA and 3.6% in CRLB. This affirmation is supported because CRLA shows higher isotherm values in this zone, than CRLB. At higher a~ values the system isooctane + isoenzyme accepts more water than the pure isoenzyme because both components are hydrated. mg
water/g
0.40
[
A
~
sample
i,ooctanel ~
0
water/g
o.40B " I t
~
sample
Isooctane~
"
"
0.20 0.00
mg
0.20 -
0.00 mg
I 0.50
Aw water/g
4.00 2.00
0.00
'
I 0.00 1.00 0.00 sample
I 0.50 Aw
I 1.00
C
'BB
Isooctane]
~
Air_
1- --~-- - I - - ' 0.00 0.50 Aw
I 1.00
Figure 2. Water sorption isotherms of several pure solid isoenzymes. Figure A: CRLA, Figure B" CRLB, Figure C: CALB. T = 25 ~ ; 100 mg solid and lml of isooctane. Finally pure isoenzyme B from Candida antarctica CALB is more hydrophile than pure isoenzymes from Candida rugosa as we can deduce from Figure 2C because CALB adsorbs more water than C. rugosa isoenzymes CRLA and CRLB. Its hydrophile nature means that the shape of both curves (Figure 2C) are similar to the case of hydrophile CCRL shown earlier. Our results agree with the very different shapes of water adsorption isotherms obtained by Valivety et al [3,6] with lipases from different sources. Unfortunately, the purification degree of the enzymes are not described in the paper and their results can only be used as a qualitative reference for us because the impurities dramatically alter the sorption isotherms (data not shown). Water sorption isotherms can be used as a tool to explore the hydrophilic/hydrophobic characteristics of chemically modified enzymes and to explain their different enzymatic activity depending on the modification degree. In Figure 3 we show, as an example, the isotherms of two chemically modified enzymes - obtained from previously dialyzed CCRL and called CRSL- with different molar rations enzyme / modifier (PNFCF-PEG): ratio 1/5 (CRSL-PNFCF- 1/5) and ratio 1/30 (CRSL-PNFCF-1/30). The modification percentage of the external lysines was 60% and 77%, respectively, equivalent to 11 and 15 modified lysines. The percentage of protein, in weight,
457 was 30% and 19%, respectively. We can observe that the less modified enzyme (CRSL-PNFCF1/5 (Figure 3A) shows a very similar water adsorption isotherm in air and sorption water isotherm in isooctane. On the contrary CRSL-PNFCF-1/30 shows a less hydrophile behavior because the isotherm of the modified enzyme in isooctane is below and further from the isotherm of the modified enzyme in air.
mg water/g
1 . 5 0 ---
sample CRSL-PNFCF-115
BB
,n?ir
A
mg water/g
1 / ~
1.50 --
1.00
1.00 --
0.50
0 . 5 0 ---
0.00
0.00
0.00
0.50 Aw
1.00
0.00
sample
B
, c.s-P.~cF.+,so 1 7' Inair / / In isooctane~
0.50
1.00
Aw
Figure 3. Water sorption isotherms of chemically modified enzymes from CRSL. 100 mg solid, 1ml of isooctane. T = 25 oc. The qualitative difference between CRSL-PNFCF-1/5 and CRSL-PNFCF-1/30 may be explained taking into the account that CRSL-PNFCF- 1/30 mg water/g solid has 19% of protein and 81% of the amphiphilic polymer but CRSL-PNFCF- 1/5 has only 70% of polymer and 30% 1.6 of protein. So, CRSL-PNFCF-1/30 is more amphiphilic . CALB silica derivative due to the presence of a large amount of polar ether 1 . 2 = in air groups (CH2-O-CH2). The practical consequence of these results is that in CRSL-PNFCF- 1/30 the protein is located in isooctane in a polar m i c r o e n v i r o n m e n t " w a t e r like" 0 . 8 microenvironment and will need a very small amount of water to be activated. From Figure 3 we can deduce that 0.4 -the "water like" microenvironment is changed by the modifier percentage: CRSL-PNFCF-1/30 > CRSL~. _ PNFCF-1/5. 0.0 1" " ! ~-- I - ' - I ' t ' I On the contrary, the less modified enzyme has the 0.0 0.2 0.4 0.6 0.8 1.0 protein in a more hydrophobic microenvironment Aw "isooctane like" than CRSL-PNFCF- 1/30 and it will have to be hydrated to be active. Figure 4. Sorption isotherm of covalent immobilized CALB on silica.T = 25 ~ mg solid and lml of isooctane. m
To generalize the study to immobilized derivatives of enzymes on solid supports, lipase B of Candida antarctica (CALB) was covalently immobilized on silica by 2,4,6-trichloro-l,3-5triazine (TCT) methodology (CALB-S). Figure 4. In the case of immobilized biocatalysts - where the protein is the minoritary component in weight - the support plays the main role in the control of the microenvironment of the enzyme. This conclusion is similar to that described for the strongly modified enzyme (CRSL-PNFCF-
458 ~/30). A practical consequence of this study is that we can predict the initial a, value of the reaction flask to achieve the maximum yield (Figure 5).
Ester Yield (%)
A
60-q
B
Ester Yield
4o t
~ 40
(%)
30 ~ r
[---~--aw:l.O[
,o l ti O
0
20~
LCRS-PNFCFoll3O 1 Ester Yield (%) --e--
,w .~.o ~
~
.w.o.o j
_,,_ --~ ...o, ,w-o-4// --'1"-- ,w.o.= [
~
I "':~ ../7/ t-+-.w:o.o, lOt ,-
~'"
40
-o ' -
1
80
Time
(h)
''
~
120
'-
T
0
'
- fI
40
'
.I .
80
Time
f
lt~_ ...,o ...o.,
30 -~F m -
.*.o.,
20~i
"w'~
/] 11- +~-
"" "" ~* ' l i
?
...o,
lO .' .
(h)
.I .
120
'.
C
CA,n-S
,
0
,
,
50
~ r I~'~
,
.... , , , ,
I00
Time (h)
,
150
,
I
200
Figure 5. Esterification yield of (R,S) 2(4-isobutyl phenyl) propionic acid (66 mM) with 1propanol (66mM), catalyzed by different enzymes. In all cases we studied the enantiomeric ratio not change by the water activity effect. A native hydrophile lipase such as CRLA gives the best results at a~ =1.0 (Figure 5A). Highly modified lipase ( such as LCRS-PNFCF 1/30, gives the best results at aw _<0.4because it is in a "water like" medium.(Figure 5B). Immobilized enzyme on silica (hydrophobic support) gives the best results for 0.8 < a, < 1.0 according to Figure 5C. REFERENCES 1. 2. 3.
C. Orrenius, T. Norin, K.Hult and G. Carrea. Tetrahedron Assymetry, 6 (1995) 3023. R.M. de la Casa, J.M. S~achez-Montero and J.V. Sinisterra.- Biotechnol.Lett 18 (1996) 13. R.H. Vaiivety, P.J. Hailing, A.D. Pellow and A.R. Macrae Biochim. Biophys. Acta 1122 (1992) 143. 4. M. Norin, J. Boutelje, E. Holmberg and K. Hult. Appl. Microbiol. Biotechnol. 28 (1988) 527. 5. M. Arroyo, J.M. Moreno and J.V. Sinisterra .J. Mol. Catal.A: Chem. 97 (1995) 195. 6. R.H. Vaiivety, P.J. Hailing, A.D. Pellow and A.R. Macrae. Biochim. Biophys. Acta 1118 (1992) 218. 7. J.M. S~chez Montero, V. Hamond, D. Thomas. and M.D. Legoy. Biochim. Biophys.Acta 1078 (1991) 345. 8. M.Rua, T. Diaz-Mourifio, V.M. Fernandez, C. Otero and A. Bailesteros. Biochim.Biophys.Acta 1156 (1993) 181. 9. J.M S~chez-Montero, A. Ferjancic-Biagini, A. Puigserver and J.V. Sinisterra in Biocatalysis in non-conventional media. J.Tramper et al. eds. Elsevier Pub pp 371.1992. 10. S. Sneyder and P.Z. Sobocinski Anal. Biochem. 64 (1975) 284. 11. J.M. Moreno and J.V. Sinisterra. J.Mol.Catal. 93 (1994) 353.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
459
T h e r m a l stability o f free and i m m o b i l i s e d Pseudomonas cepacia lipase in a q u e o u s and o r g a n i c m e d i a G. Pencreac'h and Jacques C. Baratti ERS CNRS 157, Biocatalysis in Fine Chemistry, Chemistry Department, University of Luminy, Case 901, 163 avenue de Luminy, 13288 Marseille cedex 9, France.
The thermal stability of the purified lipase from Pseudomonas cepacia was investigated in both aqueous and organic media by using the "free" lipase and the lipase immobilised on Accurel EP-100. Both preparations were more stable in heptane than in water (597 and 240-fold for "free" and immobilised preparations, respectively). The first order inactivation constants were more sensitive to temperature changes (i.e. higher activation energy) for the "flee" lipase than for the immobilised one.
1. INTRODUCTION Lipases are widely used as biocatalysts for the synthesis of useful compounds such as flavour esters or chiral building blocks to be used in fine chemistry (1). For these applications, lipases are mainly used in organic media since most substrates are insoluble in water. The use of an organic solvent leads to an increase of the thermal stability of the biocatalyst attributed to the low water content of the reaction medium. In this work we present data concerning the lipase from Pseudomonas cepacia, an enzyme commercially available and widely used for applications.
2. MATERIALS AND METHODS
2.1. Free and immobilised lipase preparations The P. cepacia lipase (lipase PS, activity on olive oil: 32,600 IU/g) was from Amano Pharmaceutical Co. Ltd (Nagoya, Japan). It was used after purification by ion exchange chromatography as previously described (2). The purified enzyme showed a single band on SDS-PAGE. The purified preparation was directly used in aqueous medium. In organic medium, a lyophilized preparation containing Bovine Serum Albumin was used as previously described (2, 3). Both preparations are refereed as "free" lipase. Microporous polypropylene powder Accurel| EP-100 (particle size 200-1000 Bin; void volume 75%; pore size 0.05-0.5 ~tm; inner surface 90 m2/g) was from Akzo Faser AG, Obernburg, Germany. The lipase was immobilized by adsorption on this support from an aqueous solution of the purified enzyme (2, 3). This preparation was referred as immobilised lipase.
460
2.2. Thermal stability In aqueous medium, the free and irnmobilised lipases were incubated in a 10 rnM ammonium acetate buffer pH 7.0 at different temperatures (50, 60, 70 and 80~ Samples were withdrawn at regular time intervals and the remaining activity was assayed in standard conditions (at 37~ on emulsified pNPC16 (p-nitrophenyl palrnitate). In organic medium, the free and immobilised lipases were incubated at 80~ in heptane. The remaining activity was assayed in standard conditions (at 37~ in organic medium. 2.3. Lipase assay in aqueous medium The hydrolytic activity was measured on emulsified p-nitrophenyl palmitate (pNPC16) according to Kordel et al (4). One volume of a 16.5 mM solution ofpNPC16 in 2-propanol was mixed just before used with 9 volumes of a 50 mM Tris-HC1 buffer pH 8.0 containing 0.4% (w/v) Triton X-100 and 0.1% (w/v) arabic gum. Then, 1.35 rnl of this mixture was pre-equilibrated at 37~ in a 1 rnl cuvette of a UV-visible spectrophotorneter (Shirnadzu UV-160A). The reaction was started by addition of 0.15 rnl of enzyme solution at an appropriate dilution in 50 mM Tris-HC1 buffer pH 8.0. The variation of the absorbance at 410 nm against a blank without enzyme was monitored for 2-5 minutes. Reaction rate was calculated from the slope of the absorbance versus time curve by using an apparent molar extinction coefficient of 12.75 106 cm2/rnole for p-nitrophenol. This value was determined from the standard solutions of pNP in the reaction mixture. One enzyme unit was the amount of protein liberating one I.tmole of p-nitrophenol per minute in the above conditions. For the immobilised enzyme preparation Triton X-100 was omitted from the incubation mixture to avoid enzyme release from the support. 2.4. Lipase assay in organic medium The activity of the lipase preparation was determined according to Pencreac'h and Baratti (5) with minor modifications. In two or three Eppendorf tubes (2 rnl) containing varied amounts of the lipase preparation, the reaction was started by addition of 1.5 rnl of n-heptane containing 50 mM p-nitrophenyl palrnitate (pNPC16). The mixture was incubated at 37~ under reciprocal agitation at 100 strokes per minute (Clifton shaker bath NES-28). After 5 to 15 rnin of reaction, the mixtures were gently centrifuged (2,200 g, 1 min) and the clear supematants were withdrawn. Then, 50-400 ~tl of the supematant were mixed with 1 rnl of Tris-HC1 buffer 10 mM pH 8.0, directly in a 1 rnl cuvette of the spectrophotometer. The p-nitrophenol (pNP) liberated was extracted by the aqueous phase. The absorbance was read at 410 nm against a blank without enzyme. When necessary, i.e. absorbance above 1, the organic sample was diluted in n-heptane before the extraction step. The reaction rate was calculated from the slope of the regression line of the curve absorbance v e r s u s the amount of biocatalyst. The molar extinction coefficient of pNP was estimated at 15.09 106 crn2/rnole by using standard solutions of pNP in n-heptane and extraction as described above. One enzyme unit was the amount of biocatalyst liberating one ttmole of p-nitrophenol per minute in the above conditions.
3. RESULTS AND DISCUSSION
3.1. Stability in aqueous medium The free and immobilised lipases were incubated at different temperatures and the remaining activity was assayed at regular time intervals. The results are shown in Fig 1 and
461
100
50oC
80
60oC
.v.,~
"~
60
=
40
.v,,~
r~
20
70oC
80~ O
~
-"~3~---~
!
!
10
20
--1
30
40
!
!
50
60
70
Time (h) Figure 1. Thermal inactivation of "flee" lipase preparation in aqueous medium.
120 100
.~
80
~
6o
"~
40
50oC
20
70oC ,
0
10
80~
~
-
!
!
!
!
|
20
30
40
50
60
70
Time (h) Figure 2. Thermal inactivation of immobilised lipase preparation in aqueous medium.
462 2. The curves were quite similar for both preparations. The deactivation process followed first order kinetics as demonstrated by linearity of the logarithmic plots of k against time (data not shown). The calculated inactivation constants are given in Table 1. At 50 and 60~ the inactivation constants were higher for the immobilised preparation while they were lower at 70 and 80~ This result suggested a higher sensitivity to temperature of the inactivation constant of the "free" lipase compared to the immobilised lipase. An Arrhenius plot of the log of inactivation constant against the reciprocal of temperature (Fig 3) gave linear curves from which activation energies of 182 and 94 kJ/mole could be derived for "free" and immobilised lipases, respectively. The higher activation energy indicated a higher sensitivity of the inactivation constant to an increase in temperature.
3.2. Stability in organic medium The free and immobilised lipases were incubated at 80~ in heptane and the remaining activity assayed. The results are shown in Fig 4. A very high thermal stability was observed since 55 and 37% of initial activity was recovered after 62 h of incubation for "free" and immobilised lipases, respectively. Higher temperatures could not be tested because the boiling point of heptane is 94~ Inactivation constants were deduced and are shown in Table 1. Both enzyme preparation showed similar values, the "free" lipase being more thermostable than the immobilised one. 3.3. Comparison of aqueous and organic media The thermostability of the purified P. cepacia lipase was greatly increased in heptane compared to water. The reduction of the inactivation constant was 970 and 242fold for "free" and immobilised lipase preparations, respectively. The reduction of the water content of the incubation mixture is the most likely explanation for this increased stability (6). Immobilisation did not increase the thermal stability either in aqueous or organic medium. In water, usually, an increased thermal stability is observed after immobilisation due to enzyme interaction with the support. This was surprisingly not the case in this study. In heptane, both "free" and immobilised preparations are insoluble. Therefore, the observed similar stability is no unexpected since stabilisation can occur in both eases by molecular interactions in the solid phase. From a practical point of view, the increased stability of lipase in organic medium greatly favours application for bioproeesses in these conditions rather than in aqueous phase. This will greatly improve the operational stability of the biocatalyst. REFERENCES 1. Macrae, A.R. and Hammond, R.C. (1985) Bioteclmol. Genetic Eng. 3, 193-217. 2. Pencreac'h, G., Leullier, M. and Baratti, J. (1997) 56, 181-189. 3. Pencreac'h, G. and Baratti, J. (1997) Appl. Mierobiol. Biotechnol., 47, 630-635. 4. Kordel, M., Hofman, B., Schomburg, D. and Schmid, R. D. (1991) J. Bacteriol. 173, 4836-4841. 5. Pencreac'h, G. and Baratti, J. (1996) Enz. Microb. Technol. 18, 417-422. 6. Zaks, A. and Klibanov, A. M. (1984) Science 224, 1249-1250.
463
2
1
0
Immobilised
@ 9 -2 "Free"
-3
e -~
!
2,80
,
,
i
i
2,85 2,90
i
i
I
2,95 3,00 3,05 3,10 3,15
1/T xl0 3 (~ Figure 3. Arrhenius plot for inactivation constants of "free" and immobilised lipase preparations in aqueous medium.
I
11o
100 ( 90-
=~
8o-
,~
70-
,,--,
60-
=1,-I
_~
"Free"
so
40-
~
r,/']
3o
Immobilised
20 10 0 0
i
i
i
i
i
i
10
20
30
40
50
60
70
Time (h) Figure 4. Thermal inactivation of "free" and immobilised lipase preparations in heptane at 80~
464 Table 1. Thermal inactivation constants of "free" and immobilised lipases in aqueous and organic media
Inactivationconstantk (ht)
Temperature Aqueous mcclium.,
50~ 60~ 70~ 80~
"Free" lipase
Immobiliscd lipase
0.035 0.32 3.1 9.7
0.19 1.2 2.5 3.9
0.010
0.016
Organic medium 80~
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
465
The Stabilization of Enzyme in Organic Solvent at Low Temperature. H y o n Joo Lee, Jin R y u n K i m and Young Je Yoo Department of Chemical Engineering, Seoul National University, Seoul 151-742, Korea
1. I N T R O D U C T I O N The stability of enzyme in organic media as well as in aqueous media has been a major problem in the commercial application of enzyme 1)'4). Although several merits in biocatalytic reaction in organic media were found 5), the inactivation of enzyme in organic media is serious in applications 6). The reasons for inactivation of enzyme in organic media have been investigated 7) and many techniques to enhance the stability of enzyme in organic media, such as protein engineering, solvent engineering, immobilization, using cosolvent, etc, have been proposed 8)-15) In this research, to increase the stability of enzyme in organic solvent, we performed the biocatalytic reaction at low temperature and reaction was performed in water - DMF (N, Ndimethyl formamide) solution (50 % v/v). Catalase was used as a model enzyme.
2. E X P E R I M E N T A L The catalase (EC 1. 11. 1.6, from bovine liver), PEG(polyethylene glycol, MW 3350) and D-sorbitol were purchased from Sigma(St. Louis, MO, U.S.). KCl(Potassium chloride) and DMF(N,N-dimethylforamide) were purchased from Junsei(Tokyo, Japan). peroxide (35%) was purchased from Showa Chemical.
Hydrogen
466 To measure the activities of catalase, 10 micro liter of 35 % hydrogen peroxide was added to stock solution. Potassium phosphate buffer (10mM KH2PO4-KOH, pH 7.0) was used for the experiments The activities were measured using residual hydrogen peroxide concentration. Those concentrations were measured by oxidation of hydrogen peroxide using potassium iodide by UV-visible spectrophotometer (UVIKON, Kontron, Swiss) and DO meter(DO-20A, TOA electronics Ltd.). A unit of activity was defined as the conversion of 1 mmol H202 during 1 minute. The properties of enzyme at low temperature in organic media were examined by Sybyl 6.25 (O/S Irix 6.2, Tripos Associates Inc.). The entry for high-resolution structure of catalase was taken from the selection of Protein Data Bank (PDB) 16) and the PDB entry was 8cat. The solvation of catalase was performed by silverware algorithm 17)ns) and the atomic charge was calculated by Gasteiger-Huckel method ~9). Energy minimization using Powel method 2~ and the molecular dynamics simulation were performed. In those cases, PBC (periodic boundary condition) was applied and Tripos force field21) was used. For the active site of catalase composed ofheme, Tyrosine 357 (Tyr 357), Histidine 62 (His 62), Asparagine 147 (Asn 147), the calculations were carried out at-25 ~ and 1 arm. A 10000-fs simulation was carried out by a time step of 1 Is. The temperature of system was in steady state at about 10000 fs. The monitored number of hydrogen-bonded pairs were minitored by Sybyl 6.25 using MOLCAD surfaces.
3. R E S U L T A N D D I S C U S S I O N Figure I shows the
2.0
activity of catalase in various
solutions.
water-DMF It
was
observed that catalase
t...... O . . . . . . . . . . . . . . . . O . . . . . . . . . . . . . . . . . . . . . . . . . . O . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . O
>,b A ~
-~
<:
(e
E
1.5
m ................ m .......................... m ............
........................ U
1.0
lost 90% of enzyme activity within 1 hour
o~ 0.5
when the ratio of DMF was more than 50 % (v/v). Due to several
0 D M F vlv % 20 D M F v lv % 50 D M F vlv %
.... V ....
80
DMF
vlv
%
Jib ...... .......... . ~ ....... 0.0
~ 0
-
~ 20
,,
~ 40
Incubation
reasons, the enzyme in
-9. - O - - - . .... BB..-.... A ....
9 60
time
i
!
80
100
(rain)
Figure 1. Change of catalase activity in water-DMF solution at 25 ~
120
467 organic
media
was
deactivated.
This
inactivation
makes
the
20 I
practical applications of the enzyme difficult.
.~ ..t2 .D
To increase the activity of
catalase
in
--.
organic
~.~
m
shows
the
catalase
activity
of
in
solution of
t
--~--
0.0
, 0
20
,
40
I
,
60
80
Incubation
various
Time
...
25~ O~
I
I
100
120
140
(hr)
Dependence of catalase activity on temperature in aqueous solution
F i g u r e 2.
While the catalase
,
0.5
aqueous
at
temperatures. activity
2
~
1.0
m
enzymatic reaction at low Figure
~:
_m-.
media, we performed the temperature.
1.5
<
in
aqueous solution at 25 ~ decreased with time, the activity of catalase in aqueous solution at 0 ~ was maintained stable during 130 hours, even though the initial activity of catalase at 0 ~ was lower than that at 25~ Enzymatic reaction at low temperature was performed in organic media. Figure 3 shows the activity of catalase in water-DMF solution (50% v/v) at various temperatures. In this figure, we could find that the activity of catalase in water-DMF solution (50% v/v) was maintained more
stable
as
temperature lower.
the
became
While
0.25
the
activity of catalase in this
solvent
decreased to 2 % of the initial activity within 1 hour at 25
~
the
activity of catalase at 25 ~ 80 %
< a)
= ""
0.15 E ~
C
0.10
0.05
was maintained of the initial
activity after 300 hours. These
0.20
system
imply
that
0.00
I
,
,
,
,
0
50
100
150
200
Incubation
time
250
(hr)
Figure 3. Dependence of catalase activity on temperature in waterDMF s o l u t i o n ( 5 0 % v / v )
300
468 catalase in organic media at low temperature is more stable than catalase in aqueous solution at 25 ~
This may be caused by the change of dielectric constant, pH, viscosity and
properties of noncovalent forces. The cost for enzymes can be reduced by reacting the substrates at the low temperature. In order to investigate the properties of catalase at -25 ~ in water-DMF solution (50% v/v), we performed the energy minimization and the molecular dynamics simulation at the same condition. From the results of the simulation, it was found that while at 25 ~ in waterDMF solution (50% v/v) only Asparagine was hydrogen-bonded with water molecules, at -25 ~ Asparagine, Tyrosine, Histidine, nitrogen atom of heme were hydrogen-bonded with water molecules or DMF molecules. It was supposed that this change of hydrogen-bond property might contribute to the stability of the enzyme at low temperature in organic media. It was also found from the simulation that water molecules were located near the active site at low temperature in aqueous solution. The radius of gyration of the active site was calculated. At -25 ~ in organic media as well as in aqueous solution, the radius of gyration of the active site was smaller than that at 25 ~
This result implied that the active site of catalase at low
temperature was more stable than that at 25 ~
From the calculation of local kinetic energy of
the active site, we found that the local kinetic energy of the active site in organic media was 247.95 kcal/mole at -25 ~
and 152.32 kcal/mole at 25~
So it was plausible that the
reactivity of the active site at -25 ~ might be higher than at 25 ~ due to its flexibility. These results suggested that the properties of noncovalent forces as well as covalent forces were changed at low temperature and these caused the changes in the stability and activity of enzyme. However, it might be disadvantageous in the aspect of energy cost to keep the operation temperature at -25 ~
In order to optimize between the stability of enzyme in organic media
and the energy cost, we performed the enzymatic reaction at 0 ~ using additives. From Table 1, it was found that sorbitol, when the concentration was higher than 1 g/L, enhanced the activity of catalase in water-DMF solution. These results are worthy of the effective stabilizing strategy for enzyme in organic media.
469 Table 1. Effect of sorbitol on catalase activity in water-DMF solution (50% v/v) at 0 ~ (112.5 hour incubation)
sorbitol(g/L)
0.001
0.01
0.1
1
10
50
activation ~)
---1b)
---1
---1
2.1
3
7
a) Activation is defined as the ration of the activity of catalase using sorbitol to that without using sorbitol b)-~l means that below 1
4. S U M M A R Y The activity of catalase in water-DMF solution (more than 50% v/v) decreased to 10 % of the activity of the native catalase. In order to enhance the stability of catalase in the water-DMF solution (50% v/v), we carried out enzymatic reaction at -25 ~
In this case, it was observed
that the activity of catalase was maintained stable during 300 hours. The stability of catalase in organic media at -25 ~ is better than in aqueous solution at 25 ~
Furthermore, though
the initial activity of catalase in aqueous solution at 25 ~ was higher than at -25 ~ stability of catalase in aqueous solution a t - 2 5 ~
was better than at 25 ~
the
In molecular
simulation, it was suggested that the properties of noncovalent forces as well as covalent forces were changed at low temperature and the catalase was maintained more stable by these reasons. To optimize between the stability of enzyme in organic media and operation cost, we performed the enzymatic reaction at 0 ~
using sorbitol as an additive. In this case, it was
found that the activity of catalase in water-DMF solution using sorbitol was higher than that without using sorbitol. Enzymatic reaction at 0 ~ using addtive can be therefore proposed as an effective stabilizing strategy for enzymatic reaction in organic media.
470 REFERENCES 1. Adlercreutz, P., Modes of Using Enzymes in Organic Media, Enzymatic Reactions in Organic Media, Chapman & Hall, 1996. 2. Bell, G., Hailing, P. J., Moore, B. D., Partridge, J., Rees, D. G. TIBTECH, 13 (1995) 468. 3. Klibanov, A. M..TIBTECH, 15 (1997) 97. 4. Yang, Z., Russell, A. J., Fundamentals of Non-Aqueous Enzymology, Enzymatic Reactions in Organic Media, Chapman & Hall, 1996 5. Dabulis, K., Klibanov, A. M. Biotech. Bioeng., 41 (1992) 566. 6. Schulze, B., Klibanov, A. M. Biotech. Bioeng., 38 (1991) 1001 7. Gorman, L. A. S., Dordick, J. S. Biotech. Bioeng., 39 (1992) 392. 8. Arnold, F. H.. TIBTECH, 8 (1990) 244. 9. Hailing, P. J. Biotech. Tech., 6 (1992) 271. 10. Douzou, P., The Study of Enzyme Reactions at Subzero Temperature, Structural and Functional Aspects of Enzyme Catalysis, Springer-Verlag, 1981. 11. Fink, A. L., Ahmed, A. I. Nature, 263 (1976) 294. 12. Boyd, S., Yamazaki, H. Biotech. Tech., 8 (1994) 123. 13. Marshall, C. J. TIBTECH, 15 (1997) 359. 14. Timacheff, S. N., Stabilization of Protein Structure by Solvent Additives, Stability of Protein Pharmaceuticals Part B, Plenum Press, 1992. 15. Triantafyllou, A. O., Wehtje, E., Adlercreutz, P., Mattiasson, B. Biotech. Bioeng., 54 (1996) 67. 16. Boberg, J., Salakoski, T., Vihinen, M. Proteins, 14 (1992) 265. 17. Mario Blanco. J. Comp. Chem., 12 (1991) 237. 18. I. Alkorta, H. O. Villar, J.J.Perez. J. Comp. Chem., 14 (1993) 620. 19. W. P. Purcel, J. A. Singer. J. Chem. Eng. Data, 12 (1967) 235. 20. M. J. D. Powel. Mathematical Programming, 12 (1977) 241. 21. M. Clark, R. D. Cramer, ,III, N. Van Opdenbosch. J. Comp. Chem., 10 (1989) 982.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
471
A comparative study of thermal inactivation o f e n z y m e s in supercritical carbon dioxide
A. Giel3aufa, T. Gamse a, E. Klingsbichel b, H. Schwab b and R. Marr a a Institut for Thermische Verfahrenstechnik und Umwelttechnik, TU Graz, Inffeldgasse 25, A-8010 Graz, Austria b Institut ~ r Biotechnologie, TU Graz, Petersgasse 12/1, A-8010 Graz, Austria I. INTRODUCTION Several studies on enzyme stabilities in supercritical carbon dioxide have been reported (mainly about lipases) since the first demonstrations that enzymes are active and stable in supercritical carbon dioxide [1,2,3]. About 20% of the total annual costs for running an enzymaticaUy catalyzed chemical process that uses supercritical carbon dioxide (SC-CO2) as the only solvent are expenses for the biocatalyst [4]. This fact explains the usefulness of comperative studies on enzyme stabilities. Some of the advances of enzymatic catalysis in SCCO2 are shown by the esterification reaction of acetic acid anhydride with different secondary alcohols [5]: The esterification rates are in almost all cases higher than in organic solvents and the enantiomeric excess values obtained in SC-CO2 are generally higher than those obtained in organic media [5]. Kasche et al. [6] realized first that depressurization steps are an important factor causing enzyme inactivation and these results have been confirmed in case of achymotrypsin [7]. Zagrobelny et al. [8] showed by fluorescence spectroscopy that indeed the pressurization step is the reason for a change of the protein conformation of trypsin. The thermal stability of enzymes should be involved in the consideration of the applicability of enzymes, because there are already several publications indicating that reaction rates are increasing with increasing temperature. Pasta et al. [9] found out, that the conversion rate of N-acetyl-L-phenylalanine catalyzed by Subtilisin Carlsberg by a temperature change from 45~ to 80~ increased. Kamat et al. [ 10,11 ] demonstrated that temperature has no significant effect on the reaction rate (on transesterifieation between methyl-methacrylate and 2-ethylhexanol with lipase from Candida cylind~acea) in the range of 40~ and 50~ Above 55~ however, the reaction rate is more temperature dependent (the reaction rate increases more than 20-fold between 40~ and 70~ [ 11 ]. Interestingly the half-life of lipase from Candida cylindracea ACKNOWLEDGEMENTS
This work is supported by a grant of the Austrian Science Fund FWF (project no. 1215I-CHE). Fluorescence spectra were recorded at the lnstitut ~r Biochemie, Karl-Franzens Universi~t Graz.
472 increased from 18 hours at 40~ to 40 hours at 55~ [11]. It was also observed that an increase of temperature from 40~ to 70~ leads to an increase of transesterification activity between triolein and ethylbehenate by lipozym IM [12]. Only in case of crude enzymes (Candida Lipase B [13] and lipase from Pseudomonas sp.[ 14]) a temperature optimum at 40~ and no further increase between 40~ and 60~ was observed. Although a lot of data on stabilities of enzymes are already published a comparison of a greater number of enzymes under identical conditions is still missing. 2. MATERIAL AND METHODS All chemicals used were purchased by Merck (Darmstadt) and were of p.a. quality. 1,2-O-Dilauryl-rac-glycero-3-glutaric acid resorufin ester (DGGR) and Thesit was supplied by Boehringer Mannheim and 2-nitrophenyl butyrate (o-NPB) by Fluka (Buchs). Lipase from Candida cyfindracea (EC 3.1. 1.3) was supplied by Sigma, lipase AY30 from Candida rugosa and PS from Pseudomonas sp. by Amano (Nagoya). Esterase EP10 from Pseudomonas marginata was obtained by Prof. Schwab (Institut ~ r Biotechnologie, TU Graz). Following assays were used for enzyme activity measurements before and atter treatment with SC-CO2 : esterase assay: 10ttl of a substrat solution (50.4 ttl o-nitrophenylbutyrate were dissolved in 549.6 ttl ethanol) was added to 9901al of an enzyme solution (10,1lag enzyme/ml in 0,1M tris(hydroxymethyl)aminomethan~Cl pH=7.0). The linear increase of absorbance after 5 minutes of incubation at h 420nm was used for determination of enzyme activity. lipase assay: 1001al of a substrat solution (6 mg 1,2-O-dilauryl-rac-glycero-3-glutaric acidresorufin ester were dissolved in 12ml of a 1:1 mixture of dioxan/thesit) was added to 900ttl of the enzyme solutions (0.1 M in KH2PO4 pH=6.8). The increase of absorbance at ~ 572nm was used to measure activities. All absorbance measurements were performed at 25~ with a Shimadzu UV160A spectrophotometer equipped with a Lauda RM6 temperature control. Fluorescence spectra of the intrinsic fluorescence of the proteins were recorded with a Perkin Elmer LS50B spectrofluorimeter using following settings of the instrument" excitation wavelength : 280 nm, emission wavelength : 300-450 nm excitation slit: 3 nm / emission slit: 3 nm, scan speed 100 nm/min. The proteins were dissolved in the same buffer solutions as used for activity measurements and the emission spectra were recorded at 25~ 3. RESULTS AND DISCUSSION Figure 1 to Figure 6 show a comparison of two enzymes concerning their stabilities in SCCO2. Esterase EP10 (a very crude preparation of this novel enzyme that has already been used for the transesterification of the secondary terpene alcohol menthol [15]) shows a rapid decrease of enzyme activity at temperatures above 75~ Even after incubation for only 1 hour at 75~ enzyme activity of esterase EPI0 decreased to a value of 87,0 + 13,5 % (Figure 1)
473
whereby a small increase to l 11,4 + 2.8 % (Figure 4) was observed in case of lipase from Candida cylindracea. The activities of the untreated enzymes were set to 100% and is shown on the ordinate. The time course of activity change at 75~ is shown in Figure 2 and Figure 5.
~ 120
~140
100
/.-----.F-.---
40
~ 40
0
25
,
;
.
35
45
55
.
.
65
.
.
0
75
85 95 105 Temperature (~ Figure 1: Thermal stability of esterase EPIO in SCCO=
25
35
45
55
65
75 85 95 105 Temperature (*C) Figure 4: Thermal stability of lipase from
Candida cylindracea
120
140
.~ 120
L
~oo
so ~8o ~4o
o
|
~
e
=: 20
2O
0
4
8
12
18
20 24 Time (hours) Figure 2: Stability of esterase EPIO at 75"C 120
~1oo "~ 80
4
8
12
16
Figure 5: Stabilityof lipase from C,and/da cyftndmcea at 75~ ~,120 ~ ~,100
20 24 Time (hours)
* |
o
~60 .1.3.
|
40
~
40
20
0 5 10 15 20 25 30 Number of pressurization/depressurization steps Figure 3: Effect of (de-)pressurization on of esterase EP10
N Oumbe5f ro pressurization/depressurization 10 15 20 25 steles 10 Figure 6: Effectof de-(pressurization)on activity of
llpase from Candidacylindracea
474 Within the first hours of treatment of esterase EP10 with SC-CO2 at 75~ there is a rapid loss of enzyme activity but lipase from CaTgtida cylindracea is stable for 24 hours. The residual activities of the enzymes after 24 hours at 75~ and 150 bar are shown in table 1. Figure 3 and Figure 6 show the effect of pressurization / depressudzation steps on enzyme activities of these two enzymes. Again lipase from Candida cylindracea is not inactivated by 30 de-/pressurization cycles. Table 1 The residual activities of four enzymes after 30 pressurization / depressurization cycles (incubation for I hour in SC-CO2 at 150 bar and 35~ before each depressurization) are shown as well as the enzyme activities after 24 hours incubation at 75~ (I lipase from Candida c~,lindracea, II lipase PS, III !i.pase AY, IV esterase EP 10). etmyme I II III IV 75~ Residual activity (%) 110 + 4.1 Pressurization/depressurization Residual activity (%) 103 + 3.2
91 + 17.2
55 + 2.2
54 + 7.6
70+ 11.8
72+ 4.6
51 • 7.5
All enzymes show no significant loss of activity after incubation for 1 hour at 150 bar in SCCO2 at temperatures below 65~ In case of all four enzymes no changes of fluorescence emission maxima (> 2nm) of the tryptophan residues were observed before and after treatment of the enzymes with SC-CO2 at temperatures below 65~ (see also table 2). Table 2 Fluorescence emission maxima of esterase EP10 and lipase from Candida cylindracea (for experimental details read section materials and methods) Temperature [~ Esterase EP 10 Lipase from Candida cvli~acea ,
untreated 35 45 55 65
335 335 335 335 334
,
339 339 339 340 341
These results indicate that no conformational change of the studied proteins occurs caused by treatment with SC-CO2. Above this temperature significant differences between the temperature stabilities of the enzymes can be found. Lipase from Candida cylindracea was incubated at the same temperatures for 1 hour in a dryer (under air-oxygen) because it was unclear by which mechanism enzyme inactivation occurs. Interestingly the enzyme is more stable in air-oxygen than in SC-CO2 (see table 3). These results indicate that enzyme
475 inactivation at temperatures > 85~ is not caused by oxidation of certain amino acid residues (due to the use of technical carbon dioxide for all experiments) because lipase from Candida cylindracea is more stable in air. Table 3 The residual activities of lipase from Candida cylindracea after treatment with SC-CO, (1 hour, 150 bar) an.d under air oxygen our, atmospheric pressure). Temperature [~ SC-COz ..... Air-oxygen 75 85 95 105
111.4 • 2.8 94.3 + 5.6 57.9• 3.0• 1.6
127.3 + 5.8 121.8 • 2.8 131.0• 1.6 94.8 + 17.8
A possible explanation for the lowest temperature stability of esterase EP10 is the low purity of the enzyme preparation. The results shown in Figure 2 fit very well to this hypothesis, because a decomposition process may cause enzyme inactivation (after about 4 hours almost all of the unstable compound is consumed and enzyme inactivation reaction is slowed down). The moderate stability of this enzyme against pressurization/depressurization can not be explained by the fact that a crude enzyme was used (figure 3), because at 35~ a decomposition process should be of less importance. By the experiments we have performed, it is difficult to speculate whether enzyme purity is an important factor for enzyme stability in SC-CO2. Further experiments are necessary to answer this question. Therefore in the near future the stabilities of preparations with different purities of one enzyme will be compared at the department. Kasche et al. [6] observed first that enzymes with S-S bridges (c~chymotrypsin,trypsin) were denatured to a lesser degree than the enzyme without cysteine (penicillin amidase). It is intended to increase the number of enzymes used for enzyme stability studies under comparative conditions to test the hypothesis if the number of disulfid-bridges is really effecting to enzyme stability in SC-CO2. REFERENCES
.
.
5. 6.
T.W. Randolph, H.W. Blanch, J.M. Prausnitz and C.R. Wilke, Biotechnol. Lett., 7 (1985) 325. D.A. Hammond, M. Karel and A.M. Klibanov, Appl.Biochem.Biotechnol., 11 (1985) 393. K. Nakamura, Y.M. Chi, Y.Yamada, and T. Yano, Chem.Eng.Commun., 45, (1986) 207. O. Aaltonen, M. Rantakyla ChemTech. 21 (1991), 240. E.Catoni, E.Cernia and C.Palocci, J.Mol.Cat. A:Chem., 105,(1996) 79. V. Kasche, R. Schlothauer and G. Brunner, Biotechnol.Lett., 10, (1988) 569.
476
.
9. 10. 11. 12. 13. 14. 15.
E Lozano, A. Avellaneda, R. Pascual and J.L.Iborra, Biotechnol.Lett., 18, (1996) 1345. J. Zagrobelny, F.V. Bright, Biotechnol. Prog., 8, (1992) 421. P. Pasta, G. Mazzola, G. Carrea and S. Riva, Biotechnol.Lett.,II,(1989) 643. S. Kamat, J.Barrera, E.J.Beckman, A.J.Russell, Biotechnol.Bioeng.,40 (1992) 158. S. Kamat, G.Critchley, E.J.Beckman, and A.J. Russell, Biotechnol.Bioeng.,46 (1995) 610. S.-H. Yoon, O.Miyawaki, K.-H.Park and K.Nakamura, J.Ferment.Bioeng., 82 (1996) 334. D.C. Steytler, P.S. Moulson, and J. Reynolds, Enzyme Microb.Technol. 13 (1991)221. E.Cemia, C.Palocci, F.Gasparrini and D. Misiti, Chem.Biochem.Eng. 8 (1994) 1. H. Michor, R.Marr, T. Gamse, T.Schilling, E.Klingsbichel and H.Schwab, Biotechnol.Lett. 18 (1996) 79.
Stability and Stabilization of Biocatalysts A. BaUesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
477
Studies on the stability o f a m i n o a c y l a s e in some organic solvents L. Boross a J. Kosfiry~ [~. Stefanovits-BhnyaP C.
Sisak b
and B. Szajhni c
aDepartment of Chemistry and Biochemistry, University for Horticulture and Food Industry, Budapest, Hungary bResearch Institute of Chemical Engineering, Pannon University of Agricultural Sciences, Veszprrm, Hungary cCOVENT Co. Inc., Budapest, Hungary
1. INTRODUCTION Aminoacylase (N-acylamino acid amidohydrolase, EC 3.5.1.14) is a hydrolase which catalyses not only hydrolysis of N-acyl-L-amino acids but their reverse synthesis in organic solvents of low water content [1-3]. Nowadays the application of reverse hydrolytic processes in non-conventional media is of increasing interest [4,5]. However, organic solvents influence not only the catalytic properties of enzymes but also their stability. Our recent results about the effects of dimethylformamide and dioxane on the stability of aminoacylase are presented in this paper.
2. MATERIALS AND METHODS 2.1. Materials Aminoacylase isolated from pig kidney with a specific activity of 2,000-3,000 U mg ~ protein was a commercial preparation (A 3010, Grade I) of SIGMA Chemical Co (St. Louis, MO, USA). N-Acetyl-DL-methionine was purchased from REANAL Factory of Laboratory Chemicals (Budapest, Hungary). N,N-dimethylformamide (DMF), 1,4-dioxane and other chemicals were commercial products of A.R. grade. DMF and dioxane were dried by distillation from phophorous pentoxide to remove the water content followed by a treatment with Molecularsieve Union Carbide Type 3 A (Fluka AG). Aminoacylase immobilized on a polyacrylarnide-type bead support (ACRYLEX C100) possessing carboxylic groups activated by water soluble carbodiimide with an activity of 3200 U g-~ dry gel was also a product of REANAL. The binding capacity of the support was 6.2+0.3 meq/g solid and particle size was 100-320 ).tm. 2.2. Assay of enzyme activity The activity of aminoacylase was assayed at 25 ~ by the ninhydrin method adapted by [6] using N-acetyl-DL-methionine as substrate. The changes in absorbance were detected at 570 nm. One unit of activity is defined as the amount of enzyme that catalyses the
478 liberation of 1 ~tmole L-methionine in 1 hour at 25 ~ and at pH 7.0. Samples containing DMF or dioxane were assayed in less than 15 see after mixing the solvent or after appropriate incubation time.
3. RESULTS AND DISCUSSION We have shown earlier [1] that DMF and dioxane inhibit aminoacylase in concentrations higher than 2-5%. Above 25-30% solvent concentration the hydrolytic activity of the enzyme is lost. The effect of these organic solvents on the stability of aminoacylase was studied both in native and immobilized form on a derivatized polyacrylamide-type bead support containing earboxyl groups (Aerylex C-100). The enzyme was mixed with aqueous organic solvent solutions and the hydrolytic activity was measured within 15 see and after longer incubation in 0.1 M potassium phosphate buffer (pH 7.0) containing 0.015 M N-aeetyl-DL-methionine (Fig. 1 and 2). We have found that in the ease of very short (15 see) incubation in aqueous media with low concentrations of either solvent, the activity of aminoaeylase is the same as that of untreated enzyme. However, during longer incubation the enzyme loses its activity and the rate of inactivation is increasing with the enhancement in the concentration of organic solvents. Because of the extensive dilution (125 times) of starting mixtures the organic solvent content of the test solutions is negligible. It was found that inactivation by DMF follows an apparent first order reaction kinetics in lower DMF concentrations but in higher concentrations the time curves of inactivation became complex (Fig. 3). According to the preliminary data of inactivation caused by dioxane it seems that the process is much more complex (Fig. 4). The chemical nature and polarity of DMF and dioxane are different. It is suggested that DMF is able to promote the dissociation of aminoaeylase dimer and the unfolding processes of dimer and monomer are parallel. In the ease of dioxane it seems that the unfolding is faster than dissociation of dimer. Probably dioxane perturbes the hydrophobic core of the protein molecule. The covalent immobilization of aminoaeylase on Aerylex C-100 increases its stability against the organic solvents mentioned above (Fig. 5). Aminoacylase preserves at least 40% of its original activity in mixtures even of 50% content of organic solvents after 20 min incubation. The rate of inactivation is faster in 50% dioxane containing mixture than in 50% DMF. The semilogarithmic representation of residual activities shows that inactivation of immobilized aminoaeylase follows a complex kinetics in both solvent mixtures composed of at least a fast and a slow phase. A detailed study of the problem is in progress in our laboratory. In an earlier study [7] it has been demonstrated that the solvent mixtures inside the gel particles and outside are the same.
ACKNOWLEDGEMENT This work is supported the Hungarian Scientific Research Fund (Grant No. T-015558). The authors wish to express their thanks to Miss S. M. Firisz and A. B6day for skillful technical assistance.
479
Figure 1. Effect of DMF concentration and incubation time on the stability of native aminoacylase in different DMF - 0.1 M phosphate buffer (pH 7.0) mixtures. 5 mg ml ~ enzyme solution containing different DMF in 0.1 M phosphate buffer (pH 7.0) were prepared and after appropriate incubation time samples were transferred (125 times dilution) into activity test solutions containing 0.015 M N-acetyl-DL-methionine in 0.1 M phosphate buffer (pH 7.0). The specific activity (3200 ~tmol h 1 mg ~ protein) of the control i.e. enzyme solution with no DMF was taken as hundred percent.
Figure 2. Effect of dioxane concentration and incubation time on the stability of native aminoacylase in different dioxane - 0.1 M phosphate buffer (pH 7.0) mixtures. 5 mg ml ~ enzyme solution containing different dioxane in 0.1 M phosphate buffer (pH 7.0) were prepared and after appropriate incubation time samples were transferred (125 times dilution) into activity test solutions containing 0.015 M N-acetyl-DL-methionine in 0.1 M phosphate buffer (pH 7.0). The specific activity (3200 gmol h 1 mg -~ protein) of the control i.e. enzyme solution with no dioxane was taken as hundred percent.
480
~'r ~,
.~, &~
3.54 I 3.52 3.5
10%DMF
3.48
v
3.46
~
~
9
20% DMF 30% DMF
3.44
3.42 0
10
20
30
40
50
Incubation time (min)
Figure 3. Plot of the logarithm of the residual enzyme activity as a function of incubation time in different DMF - 0.1 M phosphate buffer (pH 7.0) mixtures. 5 mg ml 1 enzyme solution containing different DMF in 0.1 M phosphate buffer (pH 7.0) were prepared and after appropriate incubation time samples were transferred (125 times dilution) into activity test solutions containing 0.015 M N-acetyl-DL-methionine in 0.1 M phosphate buffer (pH 7.0). The specific activity (3200 ~tmol h ~ mg ~ protein) of the control i.e. enzyme solution with no DMF was taken as hundred percent.
6)
E
3.6 3.4 3.2
0%dioxane A . ~ 5 % dioxane lO%dioxane 15%dioxane ~,~ 20%dioxane
'7 r'A 3 ~.c_. E ~ 2.8
.~, Q" 2.6 ~9 _o~
o
~
2.4 2.2 0
10
20
30
Incubation time (min)
Figure 4. Plot of the logarithm of the residual enzyme activity as a function of incubation time in different dioxane - 0.1 M phosphate buffer (pH 7.0) mixtures. 5 mg ml 1 enzyme solution containing different dioxane in 0.1 M phosphate buffer (pH 7.0) were prepared and after appropriate incubation time samples were transferred (125 times dilution) into activity test solutions containing 0.015 M N-aeetyl-DL-methionine in 0.1 M phosphate buffer (pH 7.0). The specific activity (3200 ktmol h ~ mg ~ protein) of the control i.e. with no dioxane solution was taken as hundred percent.
481
a) 100 90
o~" v
80
.>_ o r m r= "o
60
~9
(I) n,
70 50
---.~
DMF
~
Dioxane
-.---~
DMF
40
30 20
10 I
I
I
20
40
60
Incubation time (min)
b) 3.7
"o ,_
3.5
E
3.3
&
"7,
t- m A
Dioxane
E ~
._z-
2.9
~
2.7 2.5 I 0
I
I
20
40
''
I
60
Incubation time (min)
Figure 5. Effect of incubation time on the hydrolytic activity of immobilized aminoacylase in 50% DMF or dioxane containing 0.1 M phosphate buffer (pH 7.0) mixtures (a) and semilogarithmic representation (b). 10 mg solid ml "! immobilized enzyme was suspended in 50% DMF or dioxane containing 0.1 M phosphate buffer (pH 7.0) and after appropriate incubation time enzyme was separated by centrifugation and was transferred into activity test solutions containing 0.015 M N-acetyl-DL-methionine in 0.1 M phosphate buffer (pH 7.0). The activities were impressed in % of the control i.e. non-incubated solvent-phosphate buffer suspensions containing immobilized enzyme. 4. R E F E R E N C E S
1. L. Boross, J. Kos~ry, t~. Stefanovits-B~ayai, C. Sisak and B. Szajhni, J. Biotechnol., (1998) (accepted for publication). 2. J.L. Iborra, J.M. Obon, A. Manjon and S.M. Canovas, Biocatalysis, 7 (1992), 37. 3. J. Koshry, C. Sisak, B. Szajhni and L. Boross, Biocatalysis, 11 (1994) 329. 4. K. Farber and M.C. Franssen, Trends in Biotechnol., 13 (1993) 63.
482 5. F. Terradas, M. Feston-Henry, P.A. Fitzpatrick and A.M. Klibanov, J. Am. Chem. Soc. 115 (1993) 390. 6. B. Szajhni, Acta. Biochim. Biophys. Acad Sei. Hung., 15 (1980) 223. 7. C. Sisak, J. Koshry, B. Szajhni and L. Boross, Proceedings of Third International Symposium on Biochemical Engineering, Stuttgart (1995) 190.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
483
Cutinase activity and enantioselectivity in supercritical fluids Nuno Fontes, M.Concei~.o Almeida, Silvia Garcia, C61ia Peres, Jo~o Grave, M.Raquel Aires-Barros *, Cl/mdio M. Soares, Joaquim M. S. Cabral ~, Christopher D. Maycock and Susana Barreiros* Instituto de Tecnologia Quimica e Biol6gica, Universidade Nova de Lisboa, Quinta do Marquis, Apt. 127, 2780 Oeiras, Portugal Centro de Engenharia Biol6gica e Quimica, Instituto Superior T6cnico, Av. Rovisco Pais, 1096 Lisboa Codex, Portugal
We studied the performance ofFusarium solam pisi cutinase, immobilized on a zeolite, in supercritical carbon dioxide and ethylene. Water activity had a strong effect on the catalytic activity of the enzyme, unlike pressure to 300 bar. The enzyme was very selective towards one of the isomers of 1-phenylethanol, with an enantiomeric excess of virtually 100 % which did not depend on water activity, pressure or solvent. Computer modeling of the structures of transition states for the two enantiomers led to structural differences which explained the preference of the enzyme for the (R)-isomer.
1. INTRODUCTION The properties of supercritical fluids (SCF) which make them interesting solvents for extraction also make them attractive media for conducting enzymatic reactions [ 1]. The rapid growth of the market for enantiopure compounds [2] opens interesting perspectives for the use of SCF as solvents for enantioselective biotransformations. The state of hydration of an enzyme can have a pronounced effect on its activity in nonaqueous media [3,4]. The effect of pressure on enzyme activity is usually moderate in nonaqueous liquid solvents at pressures to a few hundred bar, although often larger in magnitude than in aqueous media [5-7], and can be very significant close to the critical point of a SCF [1,6,8]. Both enzyme hydration and pressure can affect enzyme enantioselectivity [9-11]. We studied the effect of enzyme hydration and pressure on the catalytic activity and enantioselectivity of immobilized cutinase from Fusarium solani pisi suspended in supercritical CO2. The influence of the medium was probed by obtaining a few data
* Author to whom correspondence should be addressed We acknowledge the support of Junta National de lnvestigacr Cientifica e Tecnol6gica (Portugal) through the contract PRAXIS 2/2.1/BIO/34/94 and the grant PBIC/C/2037/95 (C.M.S.), and of Funda~;5.o Calouste Gulbenkian through a grant (S.B.).
484 points in ethylene. The observed enantioselectivity was analyzed on the structural differences between transition states formed by the two enantiomers.
basis of
2. RESULTS AND DISCUSSION Due to the difficulty in distinguishing between the state of hydration of the enzyme molecules and that of the support in the immobilized enzyme preparation, we correlated enzymatic properties to the water activity, aw. To quantify this parameter, we built a scale of aw vs. water concentration in the solvent, in the presence of both substrates. Our results are shown in Figure 1. The fact that aw showed a linear dependence on water concentration indicated that the activity coefficient of water in the solvent mixtures was constant over the limited concentration range covered up to saturation. The solubility of water in CO2 increases significantly as pressure is raised from 80 to 250 bar at constant temperature [ 12], and this is reflected in the relative positions of the lines.
0.8 O
> 0.6 0
e~
0.4 0.2
0
0.4
0.8 1.2 water concentration/g dm-3
1.6
2
Figure 1. Water activity in the (CO2 + vinyl butyrate + 1-phenylethanol) solvent mixtures at 35 ~ as a function of water concentration. LeR line, 80 bar; right line, 250 bar. The salt hydrate pairs used and the respective aw values at 35 ~ [13] were: Na2HPO4.2/0 (0.19), Na2HPO4.7/2 (0.69), Na2HPO4.12/7 (0.90). More details of the experimental conditions given in reference 14.
Using the scales in the figure, we studied the dependence of enzyme activity on aw. Our results are shown in Figure 2. The activity of the enzyme is strongly dependent on the amount of water available to it, reaching a maximum at about aw = 0.5 and then declining, although not to zero. In water saturated ethylene at 80 bar and 15 ~ and at otherwise identical experimental conditions, the activity of cutinase was similar to that in saturated CO2. It is reasonable to conceive that an increase in temperature to 35 ~ in ethylene would
485 give rise to an increase in enzyme activity. This again supports the notion of an adverse effect of CO2 on enzyme function, whether or not the enzymes are immobilized [3,15,16]. The effect of pressure on the catalytic activity of an enzyme depends on the magnitude and the sign of the effective activation volume for the reaction, "AV~ [ 17]. Figure 2 shows a great similarity between the activity of cutinase, converted into pressure independent units by dividing by the molar volume of the solvent, at the two pressures tested, over the whole aw range. The calculation of accurate AV~ values would require more data but the results obtained so far make it reasonable to expect small in magnitude AVis for the immobilized cutinase preparation used. 100
'
80
Q C~ C3
60 Q
O
~ 4o
Q~S~
Q O
Q
"~ 2o 0
i
0
I
0.2
t
I
I
I"
0.4 0.6 water activity
l
"
I
0.8
,
1
Figure 2. Relative Vmax/K~ for the cutinase catalyzed transesterification of vinyl butyrate by 1-phenylethanol in CO2 as a function of water activity, at 3 5 ~ Full symbols, 80 bar; open symbols, 250 bar. More details of the experimental conditions and analysis given in reference 14.
The enzyme was virtually 100 % selective towards the (R)-isomer of phenylethanol, regardless of solvent, aw and pressure. To understand this, we did a computer modeling of the structures of transition states for the two enantiomers. These differed in the position of the methyl group attached to the chiral center, as seen in Figure 3. From this figure, a plausible explanation for the enzyme preference for the (R)-isomer emerges: the burial of the methyl group in the case of the (R)-isomer and the van der Waals interactions of this group with tyrosine 119 which immobilize and stabilize the transition state. A high degree of complementarity between the protein and the complex leads to a more efficient catalytic cycle. We note that the nonaqueous solvent may penetrate the active site to some extent [7,18] and could contribute to the stabilization of the above methyl group of the (S)-isomer. But apparently, these interactions are less effective from the standpoint of catalysis than those of the methyl group with protein residues in the (R)-isomer conformation.
486
Figure 3. Hypothetical transition states for transesterification of the two enantiomers of 1phenylethanol by cutinase. Details of the modeling are given in reference 14. REFERENCES 1. Kamat, S.V., Beckman, E.J., Russell, A.J. (1995) Crit. Rev. Biotechnol. 15, 41. 2. Koskinen, A.M.P. Enzymatic Reactions in Organic Media, Koskinen, A.M.P., Klibanov, A.M., Eds., Blackie Academic & Professional, 1996, p. 1. 3. Borges de Carvalho, I., Corr~a de Sampaio, T., Barreiros, S. (1996) Biotechnol. Bioeng. 1996, 49, 399-404. 4. Bell, G., Janssen, A.E.M., Hailing, P.J. (1997) Enzyme Microb. Technol. 20, 471. 5. Kim, J., Dordick, J.S. (1993) Biotechnol. Bioeng. 42, 772. 6. Fontes, N., Nogueiro, E., Elvas, A.M., Corr~a de Sampaio, T., Barreiros, S. (1998) Biochimica et Biophysica Acta 1383, 165. 7. Michels, P.C., Dordick, J.S., Clark, D.S. (1997) J. Am. Chem. Soc. 119, 9331. 8. Chaudhary, A.K., Kamat, S.V., Beckman, E.J., Nurok, D., Kleyle, R.M., Hadju, P, Russell, A.J. (1996) J. Am. Chem. Soc. 118, 12891. 9. Orrenius, C., Norin, T., Hult, K., Carrea, G. (1995) Tetrahedron: Assymetry 6, 3023. 10. Kamat, S.V., Beckman, E.J., Russell, A.J. (1993) J. Am. Chem. Soc. 115, 8845. 11. Ikushima, Y., Saito, N., Arai, M., Blanch, H.W. (1995) J. Phys. Chem. 99, 8941. 12. Chrastil, J.(1982) J. Phys. Chem. 86, 3016. 13. Hailing, P.J. (1992) Biotechnol. Tech. 6, 271. 14. Fontes, N., Almeida, M.Conceig~o, Peres, C., Garcia, S., Grave, J., Aires-Barros, M.Raquel, Soares, C.M., Cabral, J.M.S., Maycock, C.D., Barreiros, S. (1998) Ind. Eng. Chem. Res., in press. 15. Kamat, S., Critchley, G., Beckman, E.J., Russell, A.J. (1995) Biotechnol. Bioeng. 46, 610. 16. Almeida, MC., Ruivo, R., Maia, C., Freire, L., Corr~a de Sampaio, T., Barreiros, S. (1998) Enzyme Microb. Technol. 22, in press. 17. Van Eldik, R., Hubbard, C.D. Chemistry under Extreme or Non-Classical Conditions,, van Eldik, R., Hubbard, C.D., Eds., Wiley and Spektrum, 1997, p.53. 18. Parker, M.C., Moore, B.D., Blacker, A.J. (1995) Biotechnol. Bioeng. 46, 452.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
487
Effect o f pressure on e n z y m e activity in c o m p r e s s e d gases M.Concei~ao Almeida, Nuno Fontes, Eugtnia Nogueiro, Silvia Garcia, Ctlia Peres, Anttnio Silva, Manuel Carvalho and Susana Barreiros* Instituto de Tecnologia Quimica e Bioltgica, Universidade Nova de Lisboa, Quinta do Marquis, Apt. 127, 2780 Oeiras, Portugal We studied the effect of pressure to 300 bar on the catalytic activity of subtilisin Carlsberg suspended in compressed ethane, and of immobilized Candida antarctica lipase B - Novozym 435 - suspended in compressed carbon dioxide, ethane and propane. Subtilisin activity in ethane decreased with increasing pressure and this effect was more pronounced close to the critical point of the solvent, resulting in large in magnitude pressure dependent activation volumes. The catalytic activity of Novozym was hardly affected by pressure in any of the solvents.
1. INTRODUCTION High pressure may be used advantageously in many areas of biotechnology, e.g. to prolong storage time and prevent microbial contamination in foods [ 1,2], to improve protein stability at high temperature [3], to affect enzyme activity [4] and selectivity [5]. The effect of pressure on the catalytic activity of an enzyme depends on the magnitude and the sign of the effective activation volume for the reaction, AV~. AV~ values for enzymatic reactions in aqueous solvents are generally of the order of + 50 cm 3 mol ~ [6]. Comparatively larger in magnitude AVis have been obtained for biocatalytic processes in nonaqueous media, organic [7-9] and supercritical or near-critical [8,10]. In the latter case, small changes in pressure close to the critical point can have a great effect on reactions rates. Here we examined the effect of pressure to 300 bar on the catalytic activity of subtilisin Carlsberg suspended in compressed ethane, and of immobilized Candida antarctica B lipase -Novozym 435 - suspended in compressed carbon dioxide, ethane and propane.
2. RESULTS AND DISCUSSION
2.1. Subtilisin Carisberg Water may affect the catalytic performance of enzymes in nonaqueous solvents in * Author to whom correspondence should be addressed We acknowledge the support of Junta Nacional de Investigac~o Cientifica e Tecnol6gica (Portugal) through the contract PRAXIS 2/2.1/BIO/34/94 and of Funda~o Calouste Gulbenkian through a grant (s.B.).
488 many ways, e.g through its impact on enzyme dynamics [11,12], the stabilization of transition state charges [ 13], the drive for hydrolysis over synthesis. It is thus very important to quantify enzyme hydration and to refer the observed enzymatic properties to this parameter. In the case of subtilisin, we quantified enzyme hydration by means of a mass balance with the water initially present in both the enzyme and the solvent, and the water present in the solvent at water partitioning equilibrium with the enzyme. The plot of activity as a function of enzyme hydration is a bell-shaped curve for lyophilized subtilisin Carlsberg [11,14]. Comparisons of the catalytic performance of the enzyme in different solvents should be performed at a fixed point of that curve. This procedure should also be applied to a single solvent when its physical properties vary significantly with changes in experimental conditions, as is the case with compressible solvents not far from the critical point. The effects of these changes on the water sorption isotherm for the enzyme must be accounted for, so that variations in its state of hydration may be corrected. Figure 1 shows the effect of pressure on the catalytic activity of subtilisin suspended in compressed ethane. The parameter k~t/Km was calculated by taking the fraction of intact active centers to be 15 % and invariant with pressure [9]. The line represents experiments performed in the presence of higher concentrations of alcohol [8], whereas the symbols represent points obtained with alcohol concentrations about five times lower. The use of low concentrations of substrates in the latter case ensured that the reaction mixture behaved very much like the pure solvent. The direct effect of pressure on the reaction rate constant for an elementary process, k, is determined by the activation volume, AV~, which is the difference in volume between the activated and ground states [15]: [(alnk/aP)T =-AV~/RT], where P is the pressure, T the
4.5 ~
'
O
4
I
~ 3.5 O
_= 2.5
O
i
50
I
100
o
'1
o
I
150 200 Pressure/bar
o
I
250
o
300
Figure 1. Dependence of the logarithm of the catalytic activity of subtilisin for the transesterification of N-acetyl-L-phenylalanine ethyl ester by 1-propanol on pressure, in ethane at 3 5 ~ Enzyme hydration was approximately 15 %. Concentrations of 1-propanol of nearly 0.9 M (line) and 0.2 M (symbols). More details of the experimental conditions and analysis given in reference 8.
489 temperature and R the gas constant, with k based on pressure independent units. The catalytic activity of subtilisin decreases with increasing pressure, yielding positive AV#s. The variable plotted on the Y-axis of the figure was converted into pressure independent units by dividing by the molar volume of the solvent at each pressure of interest. Using the molar volume of the solvent instead of that of the mixture may be an oversimplification close to the critical point, as discussed [8]. This could artificially lower the catalytic efflciencies obtained at lower pressures, and thus lead to smaller in magnitude activation volumes. The line in Figure 1 corresponds to a constant activation volume of 139 cm 3 mol 4 [8]. The symbols in that figure lead to pressure dependent AV%, plotted in Figure 2. Such large in magnitude AV% should reflect a local density enhancement of the solvent around the solute molecules [16].
5OO !
o 400
o 300 E 200
.~ ]oo < o
:
50
',
100
i
I
150 Pressure/bar
I
I
200
250
Figure 2. Dependence of the activation volume for the subtilisin catalyzed transesterification of N-acetyl-L-phenylalanine ethyl ester by 1-propanol on pressure, in ethane at 35 ~ Concentrations of 1-propanol of nearly 0.2 M.
2.2. Novozym 435 At water partitioning equilibrium between enzyme and solvent, there is one value of the water activity, aw, which characterizes all of the phases present in the heterogeneous system. This value is related to the state of hydration of the enzyme. For immobilized enzyme preparations, it is not possible to distinguish between the hydration of the enzyme mlecules and the hydration of the support. However, as long as aw is kept constant and solvents do not differ appreciably in hydrophylicity, enzyme hydration in the various media should be the same for a given aw [17,18]. Aw may be controlled by addition of salt hydrate pairs to the reaction medium [19]. We used this technique indirectly to build a scale of aw vs. water concentration, and could show that the transesterification activity of Novozym 435 was nearly insensitive to aw over the whole aw range in compressed propane, ethane and CO2 [20]. This allowed us to neglect
490 changes in water activity brought about by changes in solvent density when studying the effect of pressure on enzyme kinetics. The results we obtained are shown in Figures 3 and 4. Because the bulk enzymatic preparation was found to suffer from internal diffusional limitations [20], we included in the present study the smaller enzyme particles. As the figures show, pressure to 300 ba~rhas a very small effect on the performance of the enzyme 0.15 m
!1
"7 =9 0.12
A r-3
!
o
Lx
0.09
E E
~ 9 0.06
:~ 0.03 9 ,,,,4 x-.-
50
100
150 200 Pressure/bar
250
;,
300
Figure 3. Dependence of the initial rates of the Novozym catalyzed transesterification of nbutyl acetate by 1-hexanol on pressure, at 35 ~ Symbols for solvents: V, carbon dioxide, A, ethane; r'i, propane. Open symbols, bulk enzymatic preparation; full symbols, 100-300 IxM diameter enzyme particles. More details of the experimental technique given in reference 20.
"7 .~
0.1
~
0.08
i
0.06
~
AA r---3
~ . r--q
.
.
A. r--q
.
A r--q
0.04 /
"=~g'=0.02
1
50
100
150 200 Pressure/bar
250
3OO
Figure 4. Dependence of the initial rates of the Novozym catalyzed transesterification of nhexyl acetate by 1-butanol on pressure, at 35 ~ Symbols for solvents as in Figure 3.
491 in all three solvents. This could be explained in part by the fact that we were never as close to the critial point of any the solvents as in the previous study with subtilisin, but should also reflect a particular characteristic of the enzyme. Figures 3 and 4 again present evidence of a lowered catalytic activity of an enzyme in C02 [21,22]. REFERENCES
1. Pothakamury, U.R., Barbosa-Chnovas, G.V., Swanson, B.G., Meyer, R.S. (1995) Chem. Eng. Prog., March, 45-53. 2. Mozhaev, V.V., Heremans, K., Frank, J., Masson, P., Balny, C. (1994) TIBTECH 12, 493-501. 3. Mozhaev, V.V., Lange, R., Kudryashova, E.V., Balny, C. (1996) Biotechnol. Bioeng. 52, 320-331. 4. Mozhaev, V.V., Heremans, K., Frank, J., Masson, P., Balny, C. (1996) Proteins 24, 8191. 5. Guthmann, O., Schwerdtfeger, R., Ricks, A., Antranikian, G., Kasche, V., Brunner, G. (1996) in High Pressure Chemical Engineering, von Rohr, P., Trepp, C., eds., Process Technology Proceedings 12, Elsevier, p. 127-131. 6. Morild, E. (1981) Adv. Prot. Chem. 34, 93-166. 7. Kim, S., Johnston, K.P. (1987) Ind. Eng. Chem. Res. 26, 1206-1213. 8. Fontes, N., Nogueiro, E., Elvas, A.M., Corr~a de Sampaio, T., Barreiros, S. (1998) Biochimica et Biophysica Acta 1383, 165. 9. Michels, P.C., Dordick, J.S., Clark, D.S. (1997) J. Am. Chem. Soc. 1997, 9331-9335. 10. Kamat, S.V., Beckman, E.J., Russell, A.J. (1995) Crit. Rev. Biotechnol. 15, 41-71. 11. Affieck, R., Xu, Z.F., Suzawa, V., Focht, K., Clark, D.S., Dordick, J.S. (1992) Proc. Natl. Acad. Sci. USA 89, 1100-1104. 12. Burke, P.A., Griffin, R.G., Klibanov, A.M. (1992) J. Biol. Chem. 267, 20057-20064. 13. Xu, Z.-F., Affleck, R., Wangikar, P., Suzawa, V., Dordick, LS., Clark, D.S. (1994) Biotechnol. Bioeng. 43, 515-520. 14. Corr~a de Sampaio, T., Melo, R.B., Moura, T.F., Michel, S., Barreiros, S. (1996) Biotechnol. Bioeng. 50, 257-264. 15. Savage, P.E., Gopalan, S., Mizan, T.I., Martino, C.J., Brock, E.E. (1995) AIChE J. 41, 1723. 16. Kim, S., Johnston, K.P. (1987) Am. Chem. Soc. Symp. Ser. 329, p.42-55. 17. McMinn, J.H., Sowa, M.J., Charnick, S.B., Paulaitis, M.E. (1993) Biopolymers 33, 1213-1224. 18. Parker, M.C., Moore, B.D., Blacker, A.J.(1995) Biotechnol. Bioeng. 46, 452-458. 19. Zacharis, E., Omar, I.C., Partridge, J., Robb, D.A., Hailing, P.J. (1997) Biotechnol. Bioeng. 55,367-374. 20. Almeida, M.C., Ruivo, R., Maia, C., Freire, L., Corr~a de Sampaio, T., Barreiros, S. (1998), Enzyme Microb. Technol. 22, in press. 21. Kamat, S., Critchley, G., Beckman, E.J., Russell, A.J. (1995) Biotechnol. Bioeng. 46, 610-620. 22. Borges de Carvalho, I., Corr~a de Sampaio, T., Barreiros, S. (1996) Biotechnol. Bioeng. 49, 399-404.
a This Page Intentionally Left Blank
Immobilized enzymes
a This Page Intentionally Left Blank
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
495
Activity and structural stability of adsorbed enzymes W. Norde and T. Zoungrana Laboratory for Physical Chemistry and Colloid Science, Wageningen Agricultural University, P.O. Box 8038, 6700 EK Wageningen, The Netherlands*)
A proteolytic enzyme, {x-chymotrypsin, and a lipolytic enzyme, cutinase, were adsorbed from aqueous solution onto sorbent surfaces of varying hydrophobicity and morphology. With both enzymes the affinity of adsorption is larger for the hydrophobic surface. Water-s01uble, flexible oligomers attached at the sorbent surface cause a reduction of the protein adsorption affinity. Circular Dichroism (CD) spectroscopy as well as Differential Scanning Calorimetry (DSC) indicate severe structural perturbations in the enzymes resulting from adsorption. The degree of structural perturbation, that is the fraction of the enzyme molecules that is perturbed by adsorption, decreases with decreasing hydrophobicity of the sorbent surface, with increasing degree of coverage of that surface by the enzyme, and with the presence of watersoluble oligomers at the sorbent surface. The specific activities of the enzymes are reduced upon adsorption, more or less following the extent of structural perturbation. When the enzyme is adsorbed its specific activity is much less sensitive to temperature variation.
1. INTRODUCTION In various biocatalytic processes, especially in the food and biomedical industries, enzymes immobilized on a (solid) matrix are employed. Immobilization allows a flow process in the bioreactor and it facilitates recovery of the enzyme. Thus, using immobilized enzymes may render the process less costly and/or more sustainable. Among the various methods of enzyme immobilization [1 ] physical adsorption may be the simplest one and it can be performed under relatively mild conditions. To use them successfully adsorbed enzymes must still be biologically active. This requirement should not be taken for granted, since upon adsorption the enzyme changes its environment which may lead to a rearrangement in the 3-dimensional protein structure. This, in turn, could very well affect the enzymatic activity. In this paper we discuss the influence of adsorption on the protein structure and on the enzymatic activity. The protein structure was studied by differential scanning calorimetry (DSC) and by circular dichroism (CD) spectroscopy. These two techniques are complementary" DSC provides information on the overall protein structural stability with respect to thermal unfolding, whereas CD probes the sub-molecular, secondary structure of the protein. *) The support from the Graduate School VLAG, The Netherlands, and from the Human Capital and Mobility program of the European Commissionis gratefully acknowledged.
496 The enzymatic activity was established as the specific activity, SA (= activity per unit mass of enzyme), - temperature profiles. Performing these types of experiments the relation between protein structure and enzymatic activity was evaluated. This was done for the enzymes in solution as well as in the adsorbed state, thereby gaining information on the effect of adsorption on the structure-function relationship of the enzymes.
2. MATERIALS 2.1. Enzymes and Substrates ct-chymotrypsin was supplied by Merck, Germany (lot no. 102307). N-acetyl tyrosine ethylester (ATEE), was purchased from Sigma. Cutinase from Aspergillus niger orizae was received from Unilever Research Laboratory, Vlaardingen, The Netherlands. Para-nitrophenyl butyrate (pNPB) was supplied by Sigma. In Table 1 some characteristics of the two proteins are given. 2.2. Sorbent materials The sorbents were supplied as colloidal particles. Teflon particles (DuPont de Nemours) consist of perfluoroalkoxyfluorocarbon resin [2]. This material does not contain UV-adsorbing double bonds or aromatic groups. The refractive index of this teflon (1.35) is comparable to that of water (1.33) so that it shows only very little scattering in the far UV. Because of the negligible UV absorption and the insignificant light scattering the teflon particles are suitable as sorbents in the investigation of the circular dichroism of adsorbed proteins. PS are negatively charged polystyrene particles prepared as described in references [3] and [4]. PS-(EO)8 are the same PS particles on which oligo- (ethylene oxide) moieties are covalently grafted [5]. Each of these oligomers consists of 8 monomers of ethylene oxide. Because the average density of (EO)8 on the PS surface is ca. one oligomer per 2.5 nm 2 [5] and the (EO)8oligomers are highly soluble in water, the PS-(EO)8 particles are sterically stabilized in an aqueous environment.
Table 1 Physical chemical characteristics of the proteins and the sorbent particles proteins
molar mass (Da)
dimensions (nm 3)
isoelectric point (pH-units)
o~-Chymotrypsin Cutinase
25200 20600
5.1 x 4.0 x 4.0 4.5 x 3.0 x 3.0
8.1 7.8
sorbents
surface morphology
water wettability
surface charge at pH 7.1
Teflon PS PS-(EO)8 Silica Aerosil OX-50 Silica Ludox HS-40
smooth smooth hairy smooth smooth
hydrophobic hydrophobic hydrophobic hydrophilic hydrophilic
negative negative negative negative negative
specific surface area (m2 g-l) 12.5 10.3 12.8 44 208
497 Aerosil OX-50 (Degussa) consists of pyrogenic non-porous silica particles. This silica was used except for the CD-measurements in which Ludox HS-40 (Du Pont de Nemours) was supplied. The silica particles of Ludox HS-40 are ultrafine; the large area/volume ratio allows a small volume fraction of silica in the samples so that interference of the circular dichroism signal from the protein by light scattering from the particles can be greatly reduced. Additional properties of the sorbent materials are included in Table 1. 3. METHODS
3.1. Adsorption Adsorption experiments were carried out at room temperature (ca. 22~ in phosphate buffer pH 7.1 and 0.01 M ionic strength. Varying amounts of protein were added to a constant amount of sorbent material so that the final volume of the mixture was 10 cm 3. The 10 cm 3tubes in which essentially no head-volume was present, were incubated for 16 hours in an endover-end rotator. Then, the samples were centrifuged and the protein concentration in the clear supematant was determined by UV absorbance at 280 nm. The amount of protein adsorbed was calculated from the difference between the protein concentrations before and after adsorption.
3.2 Desorption The protein-covered particles in the precipitate after centrifugation (see sec. 3.1) were redispersed in 10 cm 3 0.01 M phosphate buffer pH 7.1 or in 0.01 M NaCI solution, incubated for 16 hours in an end-over-end rotator and centrifuged again. The supematant was examined by UV for the presence of dilution-induced protein desorption.This procedure was executed at various elevated temperatures to check heat-induced desorption.
3.3 Circular dichroism spectroscopy CD spectra were recorded in the far UV region (185-260 nm) in a Jasco J-600 spectropolarimeter using protein concentrations ranging between 0.3 and 1.0 mg cm -3 in a 0.2 mm cuvette. From the far UV-CD spectra the secondary structure of the protein may be inferred [6]. The CD spectrum is in particular sensitive to the a-helical content of the protein: the ahelix content was estimated from the ellipticity at 222 nm. With the ultrafine silica particles (Ludox HS - 40) the adsorbed amount of protein could not be accurately determined because of incomplete separation of the particles from the solution. Hence, with these systems the influence of adsorption on the protein structure was assessed by comparing spectra of samples containing varying ratios of adsorbed and unadsorbed proteins.
3.4. Differential scanning calorimetry DSC measurements were performed in a Setaram Micro-DSC III. The sample cell contained 1-3 mg of protein and the reference cell the appropriate blank material (buffer solution or sorbent dispersion). The cells were thermally equilibrated at 20~ inside the calorimeter before heating at a rate of 0.5~ per minute up to 100~ whereafter they were cooled down to 20~ and re-heated at the same rate to 100~ again.
498
3.5. Enzymatic activity measurements Enzymatic activities of (x-chymotrypsin and cutinase at pH 7.1 were calculated from the rates of hydrolysis of ATEE and pNPB, respectively. The enzymes were incubated in a solution of ATEE (0.002 M) or pNPB (0.001 M) in 0.01 M NaCl at various temperatures.
4. RESULTS AND DISCUSSION
4.1. Adsorption isotherms The adsorption data are displayed as adsorption isotherms in Figure 1, where the amount of protein adsorbed per unit sorbent surface area, F, is plotted vs. the concentration of protein in solution after adsorption, Cp. The initial rising part of the isotherms, (dF/dcp)Cp --> 0 reflects the affinity of the protein to adsorb. Thus, with both tx-chymotrypsin and cutinase the affinity for the hydrophobic surfaces is much higher than for the hydrophilic silica surface. Adsorption saturation of 2.0-2.5 mg m -2 is compatible with a complete monolayer of more or less native protein molecules. The adsorptions of (t-chymotrypsin on teflon and on PS go beyond monolayer coverage of nativelike molecules. On these hydrophobic surfaces a second layer of protein molecules is adsorbed (possibly triggered by structural rearrangements in the molecules adsorbed in the first layer) or the protein molecules are perturbed severely in such a way that more mass can be accommodated in a single layer. Cutinase adsorption does not exceed monolayer coverage. On the silica surface adsorption saturation is relatively low and the S-shaped isotherm is characteristic of a co-operative adsorption process, that is the firstly adsorbed molecules facilitate adsorption of the subsequent ones. This would lead to two-dimensional aggregates of cutinase at the silica surface. (b) E 56 f
(a)
~
O)
E 4
o4 RE 2 I:~
E
C
3
1
0
0.2
0.4
0.6 0.8 Cp/ mg cm-3
0
0.1
0.2
I
0.3 ep / mg cm-3
01,
Figure 1. Adsorption isotherms of o~-chymotrypsin (a) and cutinase (b) for different sorbent materials. 9 silica; 9 PS-(EO)8; x PS; o teflon. T = 22~ For both proteins dilution does not lead to desorption from the hydrophobic surfaces, at least if F < 2 mg m -2. This last condition suggests that the protein more loosely attached beyond F = 2 mg m -2 is adsorbed in a second layer. In the case of hydrophilic silica the adsorption is irreversible with respect to dilution for F ~< 0.5 mg m -2. In all systems increasing the temperature does not lead to detectable desorption. The other experiments, i.e. CD spectroscopy, DSC and enzymatic activity measurements were performed under non-desorbing conditions.
499 4.2. Influence of adsorption on the three-dimensional protein structure The CD spectra of the proteins, both in solution and adsorbed on silica and teflon, are shown in Figure 2. 25000 i._ 20000 0
I
(a)
(b)
-
~--\
15000
10000 5000
i
~\
\
__.
"~ " x \ E-IO000 -5000
-15000
185
I 195
i 205
I 215
I 225
235
wavelength /
nm
245
185
195
I 205
I 215
I 225
I 235
I 245
wavelength / nm
Figure 2. Circular dichroism spectra of ot-chymotrypsin (a) and cutinase (b), in solution ( ), adsorbed (1.0 mg m -2) on teflon ( n _ m ) and incubated with silica ( m m n ) . According to the low ellipticity at 222 nm the helical content in dissolved tx-chymotrypsin is relatively small. This is in agreement with literature data [7,8]. After adsorption the spectra of o~-chymotrypsin are largely shifted, especially at the hydrophobic teflon surface. The larger effect on the protein structure at the hydrophobic teflon surface is in agreement with the adsorption isotherm data where, for that sorbent, severe structural changes in the first adsorbed layer were inferred. Moreover, at 222 nm, representative of the tx-helix content, the shifts for the hydrophobic and hydrophilic surfaces are in opposite directions. Teflon induces more o~helix in tx-chymotrypsin. Such a trend has been reported more often on changing the environment of the protein from polar (aqueous) into non-polar [2,9,10]. The spectra for the various o~-chymotrypsin-silica mixtures point to a decreased helical content. The spectra are essentially invariant with the adsorbed/unadsorbed protein ratio, suggesting that after 16 hours of incubation all the protein has been exchanged between the silica surface and the solution and that after release from the silica surface the structure remains in the conformation as it was in the adsorbed state. The CD spectrum for cutinase in solution indicates that this protein contains far more o~helix than o~-chymotrypsin. The effect of the teflon surface on the cutinase structure is relatively mild. Unlike ot-chymotrypsin, cutinase shows a (small) decrease in the amount of helix. The spectral shift for cutinase on the hydrophilic silica points to a helix reduction; it is relatively large which may be related to aggregation of the cutinase molecules on the silica surface, as discussed in see. 4.1. A decreased helical content induced by adsorption on silica has also been reported for other proteins [ 11,12]. A helix reduction in proteins at a hydrophilic surface and opposite structural responses for different proteins upon adsorption at a hydrophobic surface may be explained as follows. When a protein molecule arrives at the sorbent surface, at one side of the molecule the aqueous
500 environment is replaced by the sorbent material. Apolar parts of the protein that are buried in the interior of the dissolved molecule may become exposed to the sorbent surface where they are still shielded from contact with water. As a consequence, intramolecular hydrophobic interaction becomes less important as a factor stabilizing the globular structure of the adsorbed molecule. Because hydrophobic interaction between amino acid side groups in the protein's interior supports the formation of o~-helices, a reduction of that interaction tends to destabilize helices. Decrease of the helical content is expected to occur only if the peptide units released from the helices can form hydrogen bonds with the sorbent surface. This would be possible with oxide surfaces (e.g., glass, silica, metal oxides) or with residual water molecules remaining at the (hydrophilic) sorbent surface. Then, a decrease in the helix content may lead to an increased rotational mobility along the polypeptide chain and, hence, an increased conformational entropy of the protein. However, if the peptide units do not have the possibility to form hydrogen bonds with the sorbent surface, as is the case with hydrophobic sorbents, adsorption may induce extra intramolecular hydrogen bonding, thereby promoting the formation of secondary structures as or-helices and 13-sheets. Thus, whether adsorption on a hydrophobic surface causes an increased or a decreased ordering in the protein structure depends on the delicate balance between energetically favourable hydrogen bonds on the one hand and conformational entropy of the protein on the other. The CD spectra represent an average structure of a large ensemble of protein molecules; they do not give information about the variation in the conformational states of the protein population. That kind of information may be provided by DSC data. All transitions observed in the DSC thermograms are reflected in a single endothermic peak which indicates that the thermal unfolding of the protein, in solution as well as in the adsorbed states, occurs in a single step, characteristic for one-domain proteins. As an example, the DSC thermogram for dissolved ct-chymotrypsin is shown in Figure 3. The thermogram allows the evaluation of the denaturation enthalpy AdH as the transition peak area and the denaturation temperature Td as the temperature at half-peak area.
40
50
60 temperature (~
70
80
Figure 3. Differential scanning calorimetry of o~-chymotrypsin in solution, showing the heat-induced structural transition.
Cooling and re-heating showed that the temperature-induced structural transitions proceed irreversibly. The irreversibility is probably due to aggregation of the unfolded protein molecules. It is generally agreed [e.g. 13,14] that aggregation involves a relatively small (exothermic) enthalpy change, so that AdH essentially represents the enthalpy of the unfolding process. The DSC data, AdH and Td are summarized in Table 2.
501
Table 2 Differential scanning calorimetry between 20 ~ and 100~ of ot-chymotrypsin and cutinase in solution and adsorbed on various surfaces transition adsorbed amount (mg m -2) temperature (~ 49
ot-chymotrypsin in solution
%N
enthalpy of transition (J g-l) 20.4
100
no transition no transition no transition
tx-chymotrypsin adsorbed on teflon
0.70 1.10 1.60
ct-chymotrypsin adsorbed on PS
0.63 0.96 1.34
49 48
1.7 4.4
9 20
o~-chymotrypsin adsorbed on PS-(EO) 8
0.42 0.98 1.30
46 49 50
4.2 10.3 12.4
20 50 61
t~-chymotrypsin adsorbed on silica OX-50
0.13 0.34 0.49
50 47 48
9.0 15.0 16.9
44 74 83
53
30.0
100
cutinase in solution cutinase adsorbed on teflon
0.66 1.00 1.66
cutinase adsorbed on silica OX-50
0.10 0.14 0.24 . . . . . .
no transition
no transition no transition no transition 55 51 52
6.0 9.2 20.5 ,
,
20 31 68 ,,
For both ct-chymotrypsin and cutinase AdH is smaller when the proteins are adsorbed. However, Td is not significantly affected by adsorption. A decreased value for AdH recorded at the same Td indicates that the protein molecules in the adsorbed layer are in at least two different conformational states: a fraction is in a native-like conformation whereas the remainder has adopted a (or more) different conformation(s) that is thermo-stable up to, at least, 100~ This structural heterogeneity is expressed in the last column of Table 2, where %N (calculated from the value for AdH relative to that for the protein in solution) refers to the fraction of molecules that, below Td, are in a native-like conformation. It is evident that %N is smaller at the hydrophobic surfaces. This is in agreement with the general trend, reported in the literature [e.g. 15,16], that the extent of structural rearrangements in adsorbed proteins increases with increasing hydrophobicity of the sorbent surface. The influence of the (EOs)oligomers on the PS surface results in more adsorbed protein molecules retaining a native-like structure. Apparently, the (EOs)-moieties protect the protein molecules to some extent against
502
structural perturbation, probably by preventing intimate interaction between the protein and the PS surface. It is further interesting to note that %N increases with increasing degree of coverage o f the sorbent surface. This too is in line with the generally reported relation between the extent of structural changes and adsorbed amount [11, 17, 18], which is usually interpreted in terms of the ratio of the rates of deposition at the surface and of structural adaptation. The CD spectra for the two proteins adsorbed at teflon reveal that the structurally altered thermo-stable states (at least for T < 100~ still contain a significant amount of secondary structure. This may also be true for the perturbed states of the proteins at the other surfaces. (a)
12 -
0.3
~9
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o -~--Ro--1o
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9
40
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L%
50
60
rl
l 20
i
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l
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m
l 40
i
{
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~ >, N
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9 0
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o
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ox
~
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9
o
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9 o
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(b)
-'-9 0 . 0 2 -
9
"E
~.
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E ~' N
9
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8
A
0.1
~
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z~
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(a)
-
o
o
o
o
[] o
I
i
I
40 50 temperature / ~
....
~
, I
60
0
__Xt
20
v
ix
30
X
XI
40
X
X
o
I
o
I
50 60 temperature / ~
I
70
Figure 4. Specific enzymatic activities of ot-chymotrypsin (left) and cutinase (fight). a: A in solution, 9 adsorbed on silica; 9 adsorbed on PS-(EO)8; x adsorbed on PS; o adsorbed on teflon b: adsorbed on silica at various surface coverages (as indicated in Table 2). c: adsorbed on teflon at various surface coverages (as indicated in Table 2). For more detailed information is referrred to the text.
503
4.3. Influence of adsorption on the enzymatic activity It is of course relevant to know whether the adsorbed enzymes in the native-like state are as active as the ones in solution and, also, whether the adsorbed perturbed molecules still possess enzymatic activity. The results of the enzymatic activity measurements are given in Figure 4 where the specific activity is plotted vs. temperature. Both enzymes lose activity upon adsorption. The activity of o~-chymotrypsin at the hydrophobic surfaces of teflon and PS has disappeared almost completely. The presence of the (EO)8-oligomers at the PS surface provides some protection against adsorption-induced inactivation. Most enzymatic activity is retained at the hydrophilic silica surface. For ot-chymotrypsin the influence of the sorbent surface on the enzymatic activity closely follows the trend in %N. For cutinase the relation between the activity and the structural rearrangements is more ambiguous. First, on the silica surface, where %N = 67, the activity is remarkably low. This could be due to reduced accessibility of the enzyme's active sites caused by surface aggregation of the cutinase molecules (cf. sec. 4.1). Second, on teflon, where %N = 0, cutinase is still somewhat active. Apparently, the structural perturbation is such that it does not completely destroy the enzymatic activity. This result is not so surprising considering the relatively small shift in the CD spectrum of cutinase due to adsorption on teflon. Finally, the observation that after adsorption the activities of ct-chymotrypsin and cutinase are less sensitive to temperature variation, so that at high temperature, say 500-60 ~ C, the specific activity in the adsorbed state may exceed that in solution could be of great relevance for the application of immobilized enzymes in bioengineering.
5. CONCLUSIONS The conclusions from this study may be summarized as follows: ot-chymotrypsin and cutinase adsorb on hydrophobic and hydrophilic surfaces. The affinity of the proteins to adsorb (the strength of the protein-sorbent interaction) increases with increasing hydrophobicity of the sorbent surface. The proteins undergo conformational change s upon adsorption, that is a part of the protein population in the adsorbed layer is structurally perturbed whereas the rest remains native-like. The structural alteration at the hydrophilic silica surface involves a decrease in the helical content of the proteins, whereas at the hydrophobic teflon surface ct-chymotrypsin and cutinase show opposite trends with respect to their change in secondary structure. The opposite trends can be explained by the delicate balance between energetically favourable hydrogen bond formation and the resulting effect on the conformational entropy of the protein. The fraction of native-like adsorbed molecules increases with decreasing strength of the protein-sorbent interaction and also with increasing sorbent surface coverage. The reduction in the enzyme's specific activity is more or less correlated to the fraction of the native-like structure in the adsorbed layer: for o~-chymotrypsin this correlation is better than for cutinase. It seems that the perturbed structure of adsorbed cutinase still contains enzymatic activity. The presence of water-soluble ethylene oxide oligomers at the sorbent surface causes a decrease in the adsorption affinity of ot-chymotrypsin and, hence, a larger number of native-like molecules in the adsorbed layer, which, in turn, results in a higher specific enzymatic activity. In the adsorbed state the activities of o~chymotrypsin and cutinase are less sensitive to temperature variation.
504 REFERENCES 1.
2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.
M.H. Pinheiro, J.F. Kennedy and J.M.S. Cabral, in Interfacial Phenomena and Bioproducts, J.L. Brash and P.W. Wojciechowski (eds.), Marcel Dekker, New York, 1984, pp. 311-350. M.C.L. Maste, W. Norde and A.J.W. Visser, J. Colloid Interface Sci., 196 (1997) 224. J.W. Goodwin, J. Hearn, C.C. Ho and R.H. OttewiU, Colloid & Polymer Sci., 252 (1974) 464. J.W. Goodwin, R.H. Ottewill and R. Pelton, Colloid & Polymer Sci., 257 (1979) 61. M.C.L. Maste, A.C.P.M. van Velthoven, W. Norde and J. Lyklema, Colloids Surfaces A: Physicochem. Eng. Aspects, 83 (1994) 255. N.J. Greenfield, Anal. Biochem., 235 (1996) 1. N. Greenfield and G.D. Fasman, Biochemistry, 8 (1969) 4108. P. Manavalan and W.C. Johnson, Anal. Biochem., 167 (1987) 76. H.H.J. de Jongh and B. de Kruijff, Biochem. Biophys. Acta, 1029 (1990) 105. H. Vogel, F. J~ihnig, V. Hoffmann and J. Stiimpel, Biochim. Biophys. Acta, 733 (1983) 201. W. Norde and J.P. Favier, Colloids Surfaces, 64 (1992) 87. A. Kondo and J. Mihara, J. Colloid Interface Sci., 177 (1996) 214. S.P. Manly, K.S. Matthews and J.M. Sturtevant, Biochemistry, 24 (1985) 3842. P.L. Privalov and L.V. Medved', J. Mol. Biol., 159 (1982) 665. U. J6nsson, B. Ivarsson, I. LundstrSm and J. Berghem, J. Colloid Interface Sci., 90 (1982) 148. F. Grinnell and M.K. Feld, J. Biomed. Mater. Res., 15 (1981) 363. V. Ball, A. Bentaleb, J. Hemmerle, J.C. Voegel and P. Schaaf, Langmuir, 12 (1996) 1614. A. Kondo, F. Muramaki and K. Higashitani, Biotechnol. Bioeng. 40 (1992) 889.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
Covalent immobilization support
505
of g l u c o s e o x i d a s e o n A1PO4 a s i n o r g a n i c
F. M. Bautista, M. C. Bravo, J. M. Campelo, A. Garcia, D. Luna, J. M. Marinas and A. A. Romero. Organic Chemistry Department, Faculty of Sciences, University of Cordoba, Avda. San Alberto Magno s/n, E-14004 C6rdoba, Spain.*
A procedure was studied to obtain a covalent attachment of glucose oxidase (GOD) to the surface of amorphous A1PO4 used as support. Immobilization of the enzyme is carried out through the a-amino group of lysine residues through an aromatic Schiffs-base. The enzymatic activities of native and immobilized GOD systems were obtained in the oxidation of ~-D(+)-glucose to D-gluconic acid. In different experiments the percentage of immobilized enzyme was in the range 50-90 % while the percentage of catalytically active immobilized enzyme was always near 90 %. Moreover, GOD immobilization increased its efficiency and operational stability when repeatedly used with respect to soluble enzymes.
1. I N T R O D U C T I O N Over recent years, there has been considerable interest in the development of some methods for the covalent attachment of biologically active species to the surface of inorganic materials [1]. This blend has brought to light many new applications in the fields of chemical sensing and biocatalysis [2,3]. Immobilization on the surface of a support material has been proposed to decrease mass transfer limitations associated to several immobilization techniques, such as entrapment or adsorption in gels [4]. Thus, support material should provide a large surface area suitable for enzyme reactions. In this respect, a sol-gel material such as A1PO4, previously reported as metal support [5], could be an adequate support component for the covalent attachment of enzymes. Many functionalized linkers have been used in immobilization procedures throughout the E-NH2 groups in the external lysines of the enzyme as binding group [6,7]. However, the most common way is through glutaraldehyde [8], or some other aliphatic aldehydes [9], to obtain a Schiffs-base, In this study attempts are made to find a suitable procedure to obtain a covalent attachment of Glucose Oxidase (GOD) to an inorganic support such as A1PO4. Immobilization of the enzyme is carried "This work was subsidized by a grant from the DGICYT (PB92/0816), Mimsterio de Educacion y Cultura and from the Consejeria de Educacion y Ciencia (Junta de Andalucia), Espafia.
506 ~/I--oH+ H2N-CH2-~-~-NH2---~ #I-- NH - CH2- <
~/I--NH-CH2-~-NH2 (Step1)
~ - NH2+ HOC-~ -
CHO
~-- NH-CH2-~ ~~---NH-CH2--(~ -
N=CH-~f~- CHO
(Step2)
N=CH-~-CHO + H2N-GOD-Enzyme
~)--NH-CH2--~ -
N=CH-~ -
CH=N-GOD-E~e
(Step3)
Scheme 1. Support activation reactions and covalent immobilization of enzyme. out through the e-amino group of lysine residues to obtain an aromatic Schiffs-base that must be comparatively more stable than those usually obtained by using glutaraldehyde or other aliphatic aldehydes. Activation of the support is developed according to Scheme 1. 2. EXPERIMENTAL
2.1. Support synthesis and support functionalization Amorphous A1PO4 used as support was obtained by precipitation from the corresponding A1C13.6H20 and H3PO4 aqueous solutions by addition of ammonium hydroxide, according to a sol-gel method previously described [5,10,11]. In the present case, after filtration, washing with isopropyl alcohol and drying at 120 ~ the solid was calcined by heating at 350 ~ for 3h. The procedure for the textural and acid-baseproperties of these systems has been published elsewhere [5,10,11]. The surface area, determined by nitrogen adsorption from the B.E.T. method, was SBET-" 211, m 2 g~l. The surface acidity obtained by titration with pyridine (pKa= 5.25) was PY = 200, ~ tool gl, and with 2,6-dimethylpyridine (pKa= 6.99), DMPY = 249, p tool gl. Surface basicity obtained by titration with benzoic acid (pK~= 4.19) was BA = 352, ~ tool gl. These values were determined by a spectrophotometric method described elsewhere [11]. Activation of the A1PO4 support surface was initiated by anchoring a functionalized linker through the reaction of 4-aminobenzylamine with support surface -OH groups [12], according to step 1 in Scheme 1. The linker was obtained by a microwave heating reaction (15 rain at 380 w) of the support (20 g) and 4-aminobenzylamine (4 g) and, after that, the composite is made to react with
507
~.
40O0
V
~
tO L _. o
0
2000
..0
I
200
400
600
800
Wavelength (nm) Figure 1. Visible-Ultraviolet/Diffuse Reflectance spectra of different samples obtained in different steps in Scheme 1 (a) A1PO4 support, (13) 4aminobenzylamine on support (step 1) after microwave heating, (c) terephthaldicarboxaldehyde reaction (step 2) and (d) immobilized GOD through the e-amino group of lysine residues as indicated instep 3.
terephthaldicarboxaldehyde (step 2) by microwave heating (5 rain at 380 w). As a consequence of the high conjugation of the molecule, a yellow solid was obtained.
2.2. GOD immobilization and enzymatic activity Immobilization of GOD was carried out by thoroughly mixing from time to time, for 48 h at 4 ~ the activated solid (4 g) and Glucose Oxidase (Sigma Type X-S, 0.001 g) (~-D(+)-Glucose oxygen 1-oxidoreductase, EC 1134) from Aspergilus niger in 25 ml of phosphate buffer (KH2PO4, 0.03 M, pH 5.00). Finally, the immobilized GOD was collected by centrifugation and the resulting filtrate was separated to obtain the activity of supernatant GOD. Before the first use of immobilized GOD, the system was "equilibrated" at room temperature with 50 mL of phosphate buffer at the prefixed pH = 7.0 constant value for 24 h. Efficiency of Steps 1-3 are confirmed by Visible-Ultraviolet~iffuse Reflectance experiments shown in Figure. 1. Spectra were recorded on a UV-Visible Diffuse Reflectance Spectrophotometer Varian Carey 1E. Enzymatic activities of different soluble and immobilized GOD systems, at pH = 7.0 and different temperatures, were obtained in the oxidation of ~-D(+)-Glucose to D-gluconic acid, carried out under stirring by a magnetic bar in a batch reactor system (100 mL). With the aid of an automatic titrator, Crison, rood. micro TT 2050 TP, enzymatic activities were followed by the amount of NaOH 0.1 M necessary to neutralize liberated D-gluconic acid obtained from enzymatic glucose oxidation at the prefixed pH - 7.0 constant value. Glucose oxidase (10 rag) was
508
~
2 (c)
i4 d
.
g2 E
_
4
(a
r 7O 2 z O
>
0~ 0
10
20
30
40
5O
Time (hours) Figure 2. Catalytic activity of native (O) and immobilized enzyme ( 0 ) as well as the filtrate ( A ) obtained in the immobilization process under standard experimental conditions and different temperature and glucose substrate weight: (a) 25 ~ and glucose 0.1g, (b) 30 ~ and glucose 0.1g, (c) 30 ~ and glucose 0.025g. dissolved in 10 mL of phosphate buffer (KH2PO4, 0.03 M, pH 7.00). The reserve enzyme solution thus obtained (1 mL) was added to 50 mL of phosphate buffer containing glucose (2 ml of a 5 % (w/w) solution) to obtain reaction rates of native enzymes, rnat. Identical experimental conditions were used with filtrates of supernatant GOD, rf~, and with immobilized GOD systems, rimm. Reaction rates, r, (in p tool rain) were determined by taking the slope of the linear plot of NaOH consumption versus time. Results obtained are shown in Figs. 2-5.
509
.50 r
E 0
E
E
1.00
.40
.80
(a)
9 l,
(b)
.60
.30
.40
.20
.20
.10
! .......... I . 100 200
0
0.00
Time (hours) for successive uses
1
2
3
4
5
6
No. of uses
Figure 3. Influence of re-uses on catalytic activity of immobilized GOD under operating standard conditions at 30 ~ and pH 7.0 constant value. (a) When it is operated in the same batch reaction by successive addition of 0.025 g of glucose and (b) after filtration and solid washing in every use, and addition of 0.1 g of glucose. Hollow circle for native GOD.
3. R E S U L T S AND D I S C U S S I O N From experimental results in Fig. 2, the percentage of immobilized enzyme, Eimm, was determined by the difference between the catalytic activity of the native enzyme and the activity of the filtrate in the three different immobilization processes: rnat-
Eimm
rf~
--
(I)
x I00 rnat
The percentage of catalytically active immobilized enzyme, Eact, w a s obtained from the relation between the activities of immobilized and native enzymes: Eact =
rimm
(2)
x 100
rnat
The efficiency of the immobilized enzyme, with respect to the native one, can be obtained from the relation: Eact
Eefficiency=
Simm
rimm
X I00 --
rnat- rill
X I00
The different values obtained are shown in Table 1. Here we can see that, in
(3)
510 Table 1 Percentage of immobilized enzyme, Eimm, catalytically active immobilized enzyme, Eact, and the efficiency of immobilized respect to native enzymes, Eel, determined from the corresponding values of the catalytic activity of the native enzyme, rnat, the activity of the filtrate in the immobilization process, r~, and the activities of immobilized enzyme, r~nm, obtained from the results shown in Fig. 2. T (~
Glucose (g)
rnat (~ tool/rain)
rimm (~ tool/rain)
r~ (~ tool/rain)
Eimm (%)
Eact (%)
Eeff (%)
25
0.1
0.67
0.60
0.15
77.6
89.6
115.4
30
0.1
0.93
0.80
0.48
48.4
93.0
177.8
30
0.025
0.44
0.40
0.04
90.9
90.1
100.0
three different immobilization processes, the percentages of immobilized GOD obtained, E~m, were in the range 50-90 %. These values must correspond to the yield of step 3, in Scheme 1. In this connection, in Fig. 1 it is possible to follow by UV-Visible experiments, the changes obtained in the support surface by the effects of different reactions carried out in every step of Scheme 1, including GOD attachment by formation of aromatic Schiffs-bases with the lysine residues of enzyme. The existence of a high number of-OH surface groups in A1PO4 may be concluded from the great and very close amounts of acid sites titrated with DMPY with respect to those titrated with PY. It is known that DMPY is selectively adsorbed on BrSnsted acid sites, but not on Lewis acid sites because of a steric hindrance of two methyl groups, whereas sterically non-hindered PY is adsorbed on both BrSnsted and Lewis acid sites. Thus, the present results indicate that, in fact, all surface acid sites in A1PO4 titrated with both amines are BrSnsted ones. An extensive immobilization is not enough to obtain an appropriate behavior of the enzyme because, after immobilization, the enzyme sometimes became inactive. Among other possible causes, this can be due to the fact that the active sites of the enzyme may be involved in its attachment to the support surface. The high values of Eact, also shown in Table 1, indicate little deactivation of GOD throughout the immobilization process. In fact, the high values of Eel, point out that the GOD immobilization increases its efficiency in a variable extension with respect to the soluble enzyme, whichever the experimental conditions used. The operational stability in repeated use of immobilized GOD was also high. Thus, to determine the stability of immobilized GOD, one set of experiments was developed by operating continuously under batch conditions by successive addition of the same amount of glucose (0.025 g) under standard conditions: at 30 ~ and the prefixed pH 7.0 constant value. Results shown in Fig. 3(a) indicate that after five consecutive cycles (without separation of solids, operating in the same batch reaction by successive addition of glucose) for a week, residual activity Was 87.5 %. After two weeks continuously working and eight re-uses, residual activity was 22.5 %. When soluble GOD was used in the same way, its catalytic activity was negligible after twenty hours. The operational stability of immobilized GOD was also determined in
511
d
~D 2
~
"~,0
E ,9- - -
,
,
V
,
o
41D ~
aHim ~
I
oIB
alaB
aID ~
,
~aD
abm.lmaD
,,I
~
9
IV
I
I
:; 4
(:5
45 ~ (4th use)
50 ~ (5th use) [ ...................
"'"
s
>
0
0
i 20
55 ~ (6th use) ( ..................
""
~g
,
40
Time (hours)
0
20
40
Time (hours)
0
20
40
Time (hours)
Figure 4. Influence of the reaction temperature on the catalytic activity of native (dotted lines) and immobilized enzyme under standard reaction conditions. The used immobilized enzyme is always the same throughout along the experiment, so that on increasing the reaction temperature there is a new re-use.
successive reactions after separation and washing of solid biocatalyst. Thus, after every assay (between 2 and 4 days continuously working) the solid was easily recovered after centrifugation, washed and then reused with fresh substrate mixture. Results in Fig. 3(b) indicates that, after re-use in six cycles, including filtration after centrifugation in stirred tubes and solid washing with phosphate buffer, working continuously along 23 days, residual activity was 25%. Besides, according to the results in Fig. 4, immobilization also led to an important increase in the thermal stability of GOD. Here we can see how immobilized GOD is able to work after re-use conditions while the soluble enzyme was inactivated as soon as the reaction temperature increased. Thus, at 35 ~ while a re-used immobilized GOD is able to finish the conversion at a good reaction rate, soluble GOD is inactivated after 10 hours. At increased temperatures inactivation of soluble GOD is faster, while the same immobilized enzyme is able to continuously remain active, instead of the re-use conditions indicated. Many references can be obtained in the literature where immobilized GOD is used as biosensor [2,13]. Polymer membranes are in most cases used as GOD carriers [14] and, in the best cases, the operational stability was 2 to 4 weeks [2,13] and the immobilized enzyme 40% [14]. Comparatively, references to inorganic supports are always very reduced. In these cases, silica gel is the most common support where immobilization is carried out by entrapment [3] or by
512 using glutaraldehyde as covalent linker [15]. However, the results here presented lead us to conclude that GOD covalently immobilized on an inorganic support such as A1PO4 could be advantageously used in most described applications. 4. CONCLUSIONS From the results obtained we can conclude that due to their textural and acid-base properties, amorphous A1PO4 may be a very adequate support component for the covalent attaching of organic chains to give new hybridized composites [16]. In this connection, the linker here studied open many possibilities associated to the facility through polymerization for obtaining organic "spacers" of variable length, attached to the A1PO4 surface. In this way, tailormade organic/inorganic composites when appropriately functionalized, may incorporate a variety of organic molecules including biocatalysts and enzymes, like GOD, with a high efficiency and yield. REFERENCES 1. E. Katchalski-Katzir, Trends Biotechnol., 11 (1993) 471. 2. G.G. Guilbault, Analytical Uses of Immobilized Enzymes, Marcel Dekker, New York, 1984. 3. Y. Tatsu, K. Yamashita, M. Yamaguchi, S. Yamamura, H. Yamamoto and S. Yoshikaw, Chem. Lett., (1992) 1615. 4. A. Wojcik, J. Lobarzewski and T. Blaszczynska, J. Chem. Tech. Biotechnol., 90 (1990) 287. 5. F.M. Bautista, J.M. Campelo, A. Garcia, R. Guardefio, D. Luna, J.M. Marinas and M.C. Ordofiez, Stud. Surf. Sci. Catal., 78 (1993) 227. 6. J.V. Sinisterra and A.R. Alcantara, J. Mol. Catal., 84 (1993) 327. 7. M. Shimomura, H. Kikuchi, H. Matsumoto, T. Yamamuchi and S. Miyauchi, Polym. J., 27 (1995) 974. 8. V. Bulmus, H. Ayhan and E. Piskin, Chem. Eng. J., 65 (1997) 71. 9. R.M. Blanco, J.M. Guisan and P.J. HaUing, Biotechnol. Lett., 11 (1989) 1277. 10. F.M. Bautista, J.M. Campelo, A. Garcia, R. Guardefio, D. Luna, and J.M. Marinas, J. Mol. Catal. A. Chem., 104 (1996) 229. 11. J.M. Campelo, A. Garcia, D. Luna, and J.M. Marinas, J. Catal., 111 (1988) 106. 12. F.M. Bautista, C. Bravo, J.M. Campelo, A. Garcia, D. Luna, and J.M. Marinas, Fabrication Procedure of Activated Inorganic Solids for Covalent Immobilization of Lipases and other Enzymes, Spanish Patent No. P9601847 (1996). 13. F. Mizutani, S. Yabuki and S. Ijima, Anal. Chim. Acta, 300 (1995) 59. 14. J. Liu and Y. Chung, Angew. Makromol. Chem., 219 (1994) 101. 15. J. Jen, J. Zen, F. Cheng and G. Yang, Anal. Chim. Acta, 292 (1994) 23. 16. T. Koyano, M. Saito, Y. Miyamoto, K. Kaifu and M. Kato, Biotechnol. Prog., 12 (1996) 141.
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
513
The effect of site-specific i m m o b i l i z a t i o n o n the t h e r m a l stability of thermolysin-like neutral proteases J. Mansfeld a, G. Vriend b, B. Van den Burg ~, G. Venema c, V. G. H. Eijsinkd, and R. Ulbrich-Hofmann a ,Martin-Luther University Halle-Wittenberg, Department of Biochemistry/Biotechnology, Institute of Biotechnology, Kurt-Mothes-Strat~e 3, D-06120 Halle, Germany bBiocomputing, EMBL, Meyerhofstrat~e 1, D-69117 Heidelberg, Germany cDepartment of Genetics, Center for Biological Sciences, University of Groningen, Kerklaan 30, 9751 NN Haren, The Netherlands dDepartment of Biotechnological Sciences, Agricultural University of Norway, P. O. Box 5051,1432 As, Norway
Starting from a cysteine-free mutant of the thermolysin-like neutral protease from Bacillus stearothermophilus, cysteines were introduced into different positions on the enzyme surface by site-directed mutagenesis. The mutant enzymes were immobilized via the SH-groups to Activated Thiol-Sepharose 4B and their thermostabilities were compared to those of the soluble enzymes. The results showed that the effects of immobilization on stability strongly depend on the site of attachment. Binding of the enzyme via engineered cysteines in the critical unfolding region between residues 56 and 69 led to a considerable increase of thermal stability, whereas the immobilization via a cysteine introduced remote from the unfolding region yielded less stabilization. An extremely strong stabilization was obtained upon binding via T56C where the half-life at 75 ~ was increased by the factor of 24. 1. INTRODUCTION Several members of the bacterial genus Bacillus produce extracellular neutral proteases that resemble thermolysin, the extremely stable protease from Bacillus thermoproteolyticus. The different thermolysin-like _proteases (TLPs) are characterized by large differences in thermal stabilities [1]. TLPs have been shown to be irreversibly inactivated at elevated temperatures as a result of autolysis [2]. Previous extensive mutational studies led to the assumption that the local unfolding processes preceding autolysis, involve surface-located regions, mainly in the N-terminal domain of the enzyme [3]. In the case of the TLP from B. stearothermophilus (TLP-ste), the solvent-exposed region between residues 56 and 69 has been shown to be crucial for thermal stability [3]. These findings prompted us to use TLP-ste for probing our model of protein stabilization [4] which attributes the success of protein stabilization by immobilization to the fixation of a specific structural region of the protein molecule,
514 namely the region where the unfolding l~rocess starts. For this purpose, cysteine residues were introduced by site-directed mutagenesis into the above-mentioned critical unfolding region as well as in other regions of the protein. The various single cysteine-containing TLP-ste mutant enzymes were immobilized site-specifically via their free sulfhydryl groups, and the effects of immobilization on stability were studied. 2. RESULTS A N D DISCUSSION 2.1. Design of mutants
Suitable positions for the introduction of cysteine residues were selected by studying a three-dimensional model of TLP-ste (built by homology, using the crystal structures of thermolysin and the TLP from B. cereus, as described previously [1]; TLP-ste has 86 % sequence identity with thermolysin). Solvent exposition and accessibility to functional groups on the carrier material were prerequisites. Furthermore, mutations expected to result in significant clashes or other negative side effects were excluded. Starting from the cysteine-free mutant of TLP-ste C288L, the mutants G8C, T56C, N60C, $65C, N181C were constructed by site-directed mutagenesis and expressed successfully in B. subtilis DBl17. With the exception of the mutant N60C, expression levels were similar to wild-type TLP-ste. Wild-type and mutant enzymes had similar specific activities towards casein showing that the selected mutations had only marginal effects on the catalytic properties o f the enzyme. 2.2. Immobilization of mutant enzymes on Activated Thiol-Sepharose 4B The mutant enzymes were purified by Bacitracin-silica affinity chromatography [5] in the presence of fl-mercaptoethanolin order to orevent intermolecular disulfide formation. Immediately prior to binding to "Activated Thiol-Sepharose 4B (Amersham-Pharmacia Biotech), the mutant enzymes were subjected to gel filtration on Sephadex G25-superfine to remove salts and tg-mercaptoethanol. The ~es~llet~ot the site-specific immobilization of mutant TLP-ste enzymes are shown in
Table 1: Immobilization of TLP-ste mutants on Activated Thiol-Sepharose 4B i
Mutant
G8C T56C N60C $65C N181C C288L thermolysin
,
,w
,,
Activity of immobilized enzyme
Amount of bound protein
Relative activity
Coupling yield
[U/~]
Imp/~ carrier]
[%]
[%]
100.0 52.3 75.2 73.0 75.2
18.5 16.4 41.5 36.6 41.5
i
2.6 31.1 33.8 34.8 33.8
0.33 0.22 0.33 0.60 0.33
0.6 0.9
0.03 0.09
,,
0.3 0.6
515 In contrast to conventional immobilization strategies using the amino or carboxyl groups of proteins, the immobilization procedure used i n this study yields an uniform orientation of the enzyme molecule on the carrier material and permits the study of local binding effects on the thermal stability of the protease. Under the conditions used, nonspecific binding could be reduced to a minimum, as shown by the low coupling yield of the cysteine-free mutant C288L and thermolysin (Table 1).
2.3. Thermal stability Thermal stabilities of soluble and immobilized mutant enzymes were determined by following the time course of thermal inactivation at pH 7.5. In all cases, inactivation followed first-order kinetics. Introduction of cysteines into the TLP-ste mutant C288L strongly affected the thermal stabilities of the enzymes, varying from marginal effects to significant stabilization or destabilization effects. The haIf-lives of ttie soluble enzymes ranged from 0.9 min for N60C to 19.9 min for N181C at 75 ~ in comparison to 10.2 min for the wild-type enzyme. Intermolecular disulfide formation and, presumably, oxidation of cysteines are assumed to be the reason for the lower thermal stability of some of the mutants. The results of the comparison of thermal stabilities of soluble and immobilized mutant enzymes are given in Table 2. Table 2: Comparison of thermal stabilities of immobilized and soluble mutant enzymes at 75 ~C The enzymes were incubated at 75 oC in 0.05 M Tris buffer pH 7.5/5 mM CaC12. Aliquots removed after different time intervals were assayed for activity towards casein at 37 ~ Mutant G8C T56C N60C $65C N181C
.
Ratio of half-lives of immobilized and soluble enzymes 1.33+0.05 23.70+0.61 5.075:0.19 9.56+0.35 3.20+0.08
The binding of the enzyme via engineered cysteines in the critical unfolding region (T56C, N60C, S65C)led to a considerable increase of thermal stability (Table 2). In contrast, the immobilization via a cysteine introduced remote from the unfolding region (N181C) yielded only small stabilization. Immobilization of the protease via the introduced cysteine at position 8 did not result in significant stabilization of the enzyme (Table 2), whereas the formation of a disulfide b r i d g e between cysteines 8 and 60 stabilized the enzyme remarkably [6]. This confirms that the region around residue 60 is crucial for stability. Anextremely strong stabilization of the enzyme by immobilization was observed with the T56C mutant (by the factor 24 in comparison to 5 - 10 for the other positions tested in the critical region ) . By computer modelling, this effect could be explained by additional favorable interactions of one of the carboxyl groups in the spacer molecule of the carrier material with the Ca2*-binding site 3 of the enzyme Previous studies had shown that this Ca2*-binding site is one of the determinan'ts of thermal stability of TLP-ste [7, 8].
516 The results of these site-specific immobilization studies, using genetically introduced cysteines at predefined locations into TLP-ste, show that the attachment site at the protein surface is significant for the success of stabilization by immobilization.
Acknowledgements This work was supported by a research grant from the Leopoldina, the Deutsche Akademie der Naturforscher, and by the Deutsche Forschungsgemeinschaft, Bonn.
REFERENCES [1] Vriend, G. and Eijsink, V. G. H. (1993) J. Computer-Aid. Mol. Design 7, 367396 [2] VanBiochem.den j.Burg'272,B.~o,_~,E~sink' V. G. H., Stulp, B. K., and Venema, G. (1990) [3] Eijsink, V. G. H., Veltman, O. R., Aukema, W., Vriend, G., and Venema, G. (1995). Nature Struct. Biol. 2, 374 - 379 [4] Ulbrich-Hofmann, R., Golbik, R., and Damerau, W. (1993) Stability and Stabilization of Enzymes (van den Tweel, W. J. J., Harder, A., Buitelaar, R., eds.) 497- 504, Elsevier, London [5] Van den Burg, B., Eijsink, V. G. H., Stulp, B. K., and Venema, G. (1989) J. Biochem. Biophys. Methods 18, 202- 220 [6] Mansfeld, J., Vriend, G., Dijkstra, B. W., Veltman, O. R., Van den Burg, B., Venema, G., Ulbrich-Hofmann, R., and Eijsink, V. G. H. (1997) J. Biol. Chem. 272, 11152-11156 [7] Dahlquist, F. W., Long, J. W., and Bigbee, W. L. (1976) Biochemistry, 15, 1103 1111 [8] Veltman, O. R., Vriend, G., Hardy, F., Mansfeld, J., Van den Burg, B., Venema, G., and Eijsink, V. G. H. (1997) FEBS Lett. 405, 241 - 244
Stability and Stabilization of Biocatalysts A. Ballesteros, F.J. Plou, J.L. Iborra and P.J. Hailing (Editors) 9 1998 Elsevier Science B.V. All rights reserved.
517
I m m o b i l i z a t i o n o f h y d a n t o i n cleaving e n z y m e s f r o m A r t h r o b a c t e r a u r e s c e n s D S M 3747 - Effect o f the coupling m e t h o d on the stability o f the L-Ncarbamoylase M. Pietzsch ~ H. Oberreuter, B. Petrovska, K. Ragnitz and C. Syldatk Institute of Biochemical Engineering, University of Stuttgart,Allmandring 31, D-70569 Stuttgart, Germany
Different coupling methods were tested for the immobilization of the N-carbamoyl-Laminoacid amidohydrolase (L-N-carbamoylase) partially purified from Arthrobacter aurescens DSM 3747. The operational stability of the immobilized biocatalyst was measured using both consecutive batch reactions and continuously operated fixed bed reactors, while the stability of the flee L-N-carbamoylase was investigated using an enzyme membrane reactor. The long term stability of the enzyme was markedly enhanced by all immobilization methods and carriers tested. In consecutive batch reactions the operational stability remained relatively low and significant differences in biocatalysts stability were not observed between the different immobilization methods used. In contrast there were significant differences in the stability when the biotransformations were carded out using fixed bed reactors. As a result of this comparison the determination of the operational stability of the air-sensitive L-Ncarbamoylase on a batch-to-batch basis seems not to be useful. 1. INTRODUCTION Optically pure D- and L-amino acids can be produced with a theoretical yield of 100% and with high enantioselectivities using the "hydantoinase method" [ 1]. The instability of most of the enzymes involved in the hydrolysis of D,L-5-monosubstituted hydantoins and