Carbohydrate Bioengineering

CARBOHYDRATE BIOENGINEERING

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 Ottova, 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 (Alberghine et al., Editors) Volume 10 Carbohydrate Bioengineering (Petersen, Svensson and Pedersen)

Progress in Biotechnology 10

CARBOHYDRATE BIOENGINEERING Proceedings o f an International Conference Elsinore, Denmark, April 23-26, 1995

Edited by S t e f f e n B. P e t e r s e n

SINTEF UNIMED, MR-Center, N-7034 Trondheim, Norway Birte Svensson

Gamble Carlsberg vej 10, DK-2500 Valby, Denmark Sven Pedersen

NovoNordisk A/S, Novo Alle, DK-2880 Bagsvaerd, Denmark

ELSEVIER Amsterdam

- Lausanne

- New

York - Oxford - Shannon

- Tokyo

1995

Published by: Elsevier Science B.V. P.O. Box 211 1000 AE Amsterdam The Netherlands

ISBN 0-444-82223-2 @1995 Elsevier Science B.V. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher, Elsevier Science B.V., Permissions Department, P.O. Box 521, 1000 AM Amsterdam, The Netherlands. No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, the Publisher recommends that independent verification of diagnoses and drug dosages should be made. Special regulations for readers in the U S A - This publication has been registered with the Copyright Clearance Center Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01293, USA. Information can be obtained from the CCC about conditions under which photocopies of parts of this publication may be made in the USA. All other copyright questions, including photocopying outside the USA should be referred to the copyright owner, Elsevier Science B.V., unless otherwise specified. This book is printed on acid-free paper. Printed in the Netherlands

PREFACE The rapid development in glycobiology and glycotechnology has resulted in an enormous increase in our knowledge on the structure, conversion, and application of carbohydrates in industry and medicine. The Carbohydrate Bioengineering Meeting held at the LO-School in Elsinore, Denmark, April 23-26, 1995, gathered 230 scientists, mostly from Europe, with interest in carbohydrate analysis and structure; carbohydrates in medicine and glycopathology; structure, function, application, and protein engineering of carbohydrate active enzymes; oligo- and polysaccharides of industrial interest; and production of carbohydrate containing new materials. The meeting provided a forum where highly distinguished researchers presented their latest results. The strong representation of participants in the EU Biotechnology Programme on Carbohydrate Active Enzymes illustrated a great need for this type of meeting which provides a stimulating atmosphere where established scientists and students in academia and industry can get together. The contributions, both from invited and selected speakers as well as from the poster presentations spanned the broad field in a continuous manner. The Proceedings chapters 1 - 4 address glycoconjugates as modulatory and recognition molecules, structure determination using NMR and mass spectrometry, and microdialysis-chip enzyme-based sensors. Carbohydrate active enzymes turned out to be a predominant topic. Chapters 5 - 8 describe different bacterial enzymes involved either in carbohydrate metabolism, or with potential application in bio-processing of special polysaccharides including at elevated temperatures. Details on the tools utilized in analysis of mechanism of carbohydrases and carbohydrateprotein interactions are discussed in chapters 9 - 12 which include active site mutations coupled with crystal structures and synthetic substrate analogue interactions. Protein engineering of specificity and stability of two industrially important starch-degrading enzymes are described in chapters 13 and 14. The significance and engineering of electrostatics in carbohydrate active enzymes, and the role and engineering of N-linked sugar moieties in protein stability are demonstrated in chapters 15 and 16. A very timely up-date on ceUulolytic enzymes covers in chapters 17 - 21 the new three-dimensional structures and binding domains for biotechnological applications. The description in chapters 22 and 23 of transgenic plants for understanding and manipulation of starch biosynthesis and of the expression of cellulases in transgenic animals both have wide perspective in nutrition and related sciences. Examples on prospects for industrial synthesis of polysaccharides using enzymes not acting on sugar nucleotides, and the potential and details of the complex mechanism of such enzymes, are given in chapters 24 and 25. Furthermore, industrial applications using monocomponent plant cell-wall polysaccharide hydrolases, synthesis of fatty acid-carbohydrate ester surfactants,

vi and the utilization of bacteria in commercial production of sugars derivatives are described in chapters 26- 28. Any scientific meeting is benefitting from economical support. In our case we would like to express our special gratitude for the generous support from the following sponsors: Biosym

Technologies, Boehringer DAKO, Dansk Metal, European Commission/DG XII, European Congress of Biotechnology 5, Nordic Fund for Technology and Industrial Development, Novo Nordisk A/S, Nutek, Pharmacia, Pharmacia Biotech, Radiometer, Silicon Graphics and SNF. Finally, at this point we also wish to express our warm thanks to the participants for their contribution and for making a most exciting and fruitful meeting. A very special thank you to all authors for the effort they have made in the preparation of the manuscripts. We also greatly acknowledge the professional assistance of Ms. Mona K. Eidem in the preparation of this book.

Copenhagen, September 27, 1995

Sven Pedersen, Steffen B. Petersen and Birte Svensson, Editors

This Page Intentionally Left Blank

xi

TABLE OFCONTENTS

Chapter 1: Glycans of glycoconjugates as modulatory and recognition molecules

N. Sharon

1

Chapter 2: NMR studies of the structure and dynamics of carbohydrates in aqueous solution H . van Halbeek and S. Sheng

I5

Chapter 3: Linkage analysis by mass spectrometry of chemically modified oligo-saccharides from glycosphingolipids and glycoproteins B. Nilsson

29

Chapter 4: Development of a novel enzyme based glucose sensor F. Spener, R . Steinkuhl, C . Dumschat, H. Hinkers, K . Cammann and M. Knoll

49

Chapter 5: Carbohydrate binding at the active site of Escherichia coli maltodextrin phosphorylase P. Drueckes, D. Palm and R. Schinrel

59

Chapter 6: The chitinolytic system of Streptomyces olivaceoviridis

H. Schrempf Chapter 7: Properties and production of the P-glycosidase from the thermophilic Archaeon Sulfolobus solfataricusexpressed in mesophilic hosts M. Moracci, L. Capalbo, M. De Rosa, R. La Montagna, A. Morana, R. Nucci, M. Ciaramella and M. Rossi

71

77

xii Chapter 8: Contribution of subsites to catalysis and specificity in the extended binding cleft of Bacillus 1,3-1,4-13-D-glucan 4-glucanohydrolases A. Planas and C. Malet Chapter 9: Probing of glycosidase active sites through labeling, mutagenesis and kinetic studies S.G. Withers

85

97

Chapter 10: Thiooligosaccharides: toys or tools for the studies of glycanases H. Driguez

Chapter 11: Mutational analysis of catalytic mechanism and specificity in amylolytic enzymes B. Svensson, T.P. Frandsen, I. Matsui, N. Juge, H.-P. Fierobe, B. Stoffer and K.W Rodenburg

113

125

Chapter 12: The structure and function relationship of Schizophyllum commune xylanase A Mdr Bray and AJ. Clarke

147

Chapter 13: Protein engineering of cyclodextrin glycosyltransferase from Bacillus circulans strain 251 L. Dijkhuzien, D. Penninga, H.I. Rozeboom, B. Strokopytov and B.W. Dijkstra

165

Chapter 14: Oxidation stable amylases for detergents T.V. Borchert, S.F. Lassen, A. Svendsen and H.B. Frantzen

175.

Chapter 15: Electrostatic studies of carbohydrate active enzymes A. Baptista, T. Brautaseth, F. DrablOs, P. Martel, S. Valla and S.B. Petersen

181

Chapter 16: Effects of glycosylation on protein folding, stability and solubility. Studies of chemically modified or engineered plant and fungal peroxidases K.G. Welinder and J.W. Tams

205

xiii Chapter 17: Modes of action of two Trichoderma reesei cellobiohydrolases T.T. Teeri, A. Koivula, M. Linder, T. Reinikainen, L. Ruohonen, M. Srisodsuk, M. Claeyssens and T.4. Jones

211

Chapter 18: Structural studies on fungal endoglucanases from Humicola insolens G.J. Davies and M. Schiilein

225

Chapter 19: The catalytic domain of endoglucanase A from Clostridium cellulolyticum belonging to family 5: an 0r enzyme V. Ducros, M. Czjzek, A. Belaich, C. Gaudin and R. Haser

239

Chapter 20: Celluosome domains for novel biotechnological application E.A. Bayer, E. Morag, M. Wilchek, R. Lamed. S. Yaron and Y. Shoham

251

Chapter 21: Interactions of cellulases from Cellulomonasfimi with cellulose N. Din, J.B. Coutinho, N.R. Gilkes, E. Jervis, D.G. Kilburn, R.C. Miller Jr., E. Ong, P. Tomme and R.4.J. Warren

261

Chapter 22: Transgenic plants as a tool to understand starch biosynthesis J. K o ~ n n , G. Abel, V. Biittcher, E. Duwenig, M. Emmermann, R. Lorberth, F. Springer, I. Virgin, T. Welsh and l. Willmitzer

271

Chapter 23: Targeted expression of microbial cellulases in transgenic animals S. Ali, J. Hall, K.L. Soole, C.M.G.4. Fontes, G.P. Hazlewood, B.H. Hirst and Hzl. Gilbert

279

Chapter 24: Mechanism and action of glucansucrases J.F. Robyt

295

Chapter 25: Studies of recombinant amylosucrase M. Remaud-Simeon, F. Albaret, B. Canard, I. Varlet, P. Colonna, R.M. Willemot and P. Monsan

313

xiv Chapter 26: Application of cloned monocomponent carbohydrases for modification of plant materials L.V. Kofod, T.E. Mathiasen, H.P. Heldt-Hansen and H. DalbOge

321

Chapter 27: Fatty acid esters of ethyl glucoside, a unique class of surfactants O. Andresen and O. Kirk

343

Chapter 28: A wide range of carbohydrate modifications by a single microorganism: leuconostoc mesenteroides W. Soetaert, D. Schwengers, K. Buchholz and E.I. Vandamme

351

S.B. Petersen, B. Svensson, and S. Pedersen (Eds), CarbohydrateBioengineering 9 Elsevier Science B.V. All rights reserved.

Glycans of glycoconjugates as modulatory and recognition molecules Nathan Sharon Department of Membrane Research and Biophysics, The Weizmann Institute of Science, Rehovot 7 6100, Israel

Abstract

Glycoproteins and glycolipids are the most common classes of glycoconjugates. Their glycans are structurally diverse, although overlapping to some extent. The functions of these glycans are mostly unknown, but from the information available it is clear that they often modulate the physicochemical properties of the proteins to which they are attached, and sometimes also their biological activities. In addition, there is increasing evidence that many of these glycans, whether protein or lipid bound, serve as recognition determinants in moleculecell and cell-cell interactions. The knowledge accrued is having a marked impact on the biotechnological production of therapeutically useful glycoproteins and provides a strong impetus for the development of novel types of drugs for a wide range of diseases, such as microbial infections, inflammation and cancer.

1. INTRODUCTION Living organisms produce a bewildering array of glycans, many of them in the form of glycoproteins or glycolipids. These generally go under the term glycoconjugates or complex carbohydrates. The question of the function of these glycans is currently attracting increasing interest. Thus, oligosaccharides attached to proteins often modulate the physicochemical and biological properties of the latter. Furthermore, changes in glycan structure accompany many normal and pathological processes, from differentiation and development to cancer. In some cases, specific carbohydrates on cancer cells have been correlated with the tumor grade, metastatic potential and prognosis, raising the possibility that glycans may serve as markers for diagnostic purposes and as targets for drugs. Developments in genetic engineering have made possible the biotechnological production of pharmaceutically useful glycoproteins, both for research purposes and for therapeutic use. Last, but not least, is the prospect of creating carbohydrate-based drugs for the treatment of many diseases, from microbial infections to cancer. Findings made prior to the 1970's have given some indications that carbohydrates may have functions other than as structural materials and energy source [1]. In work carried out in the 1940's and 1950's it has been shown that sialic acid is the receptor for influenza virus, that sugars on the surface of enteric bacteria serve as phage receptors and that such compounds are determinants of ABO blood type specificity. The sixties witnessed the demonstration that

malignant cells have often a different sugar phenotype than their normal counterpart, and that sugars determine the lifetime of glycoproteins in the circulation. These findings were largely ignored by biochemists and biologists alike. During the last quarter of the century the situation has changed, in the beginning rather slowly, then at a steadily accelerating rate, and our knowledge of glycan structures and their functions has increased greatly. These developments have not escaped the attention of the popular press, as reflected in the article entitled "Sugar coated truths" (The Economist, September 24th, 1994): "Researchers have been bewitched by the elegance of nucleic acids and proteins. Now they are opening their eyes to the subtleties of sugars". Here I present a survey of the role of glycans in modulating the properties of the proteins to which they are attached and of their function as recognition determinants. The latter function is shared by membrane bound glycolipids, which is not surprising, in view of the identity of many glycan structures found in both types of glycoconjugate. These subjects have been extensively dealt with in two recent reviews [2,3], in which references to earlier literature can be found.

2. METHODOLOGY Different approaches are being employed in the quest to unravel the role of glycoconjugate glycans. They include modification of the glycans by purified glycosidases or transferases, use of inhibitors of glycosylation or glycoprotein processing and of cell mutants with known defects in glycosylation, and more recently, techniques of molecular genetics. Recombinant glycoproteins can be expressed in heterologous cells or organisms, resulting in different patterns of glycosylation. An extreme example is that of bacteria (e.g., E . coli) which produce completely non-glycosylated proteins. Nucleotide-directed mutagenesis can be applied to modify glycosylation sites on proteins so that they will no longer serve for glycan attachment. With N-glycoproteins, where the carbohydrate is linked to the Asn-X-Ser/Thr sequon, modification of either the first or third amino acid will abolish glycosylation at this site. In glycoproteins with more than one carbohydrate unit, whether N-, O- or both, the sites can be systematically eliminated in various combinations, to form a panel of mutants in which the roles of each carbohydrate chain can be assessed. Although very useful, these and other in vitro methods rarely provide answers to questions concerning the biological roles of carbohydrates in the intact, multicellular organism. Important information in this respect can be derived from investigations of congenital disorders involving genetic defects in protein glycosylation. A well-studied case is that of I-cell disease, a lysosomal storage disease, resulting from the inability to synthesize mannose-6-phosphate (due to a deficiency in N-acetylglucosaminyl phosphotransferase) [4]. Other examples are HEMPAS, a rare form of genetic anemia in humans [5] or the carbohydrate deficient glycoprotein (CDG) syndromes I and II, a set of multisystemic diseases with major nervous system involvement, all of which are due to deficiencies in various steps of N-glycosylation [6,6a]. Last but not least, the use of transgenic animals, still in its infancy, opens up new avenues for the study of glycan function in the intact organism [7]. Carbohydrate structures can now be manipulated by the expression of degradative enzymes that act on a particular carbohydrate in a particular linkage, of transferases that attach monosaccharides to glycoproteins or glycolipids, or of other enzymes, such as those that act on monosaccharide

substituents (e.g. O-acetyl). These structures may also be modified by the complete elimination of a particular glycosyltransferase ("knock-out" experiments). For example, insight into the role of complex or hybrid N-linked glycans in the development of embryos was recently obtained in experiments with transgenic mice, from which the gene for GlcNAc-T I has been removed, using homologous recombination [8,9]. Embryos of such mice exhibited an overall stunting of growth and lethality at around day 10 of gestation. These findings show that complex N-linked chains are not required during early development, but are necessary later, for the completion of morphogenesis.

3. MODULATION OF PHYSICOCHEMICAL PROPERTIES Quite frequently, the carbohydrate groups affect the physical properties of the protein to which they are attached (Table 1). In general, the effects are more pronounced the higher the carbohydrate content of the glycoprotein. The negative charges of sialic acid residues and sulfate groups change the solubility and modify the conformation of glycoproteins, as well as the adhesive properties of cells. These effects are of particular importance for the function of the highly glycosylated mucins which may carry sialic acid-containing oligosaccharides on as many as one third of their amino acids. As a result, they assume rigid, rod-like structures that may reach a length of several hundred nanometers. Mucin regions are often found on cell surface receptors and it is thought that the role of the rigid structures is to extend the functional domains away from the cell surface. Since the surface area of the glycans is quite significant when compared to that of the peptide moiety, they may, in addition, influence other properties of proteins, such as heat stability and susceptibility to proteolysis. These properties are of special interest to industries producing commercially used enzymes, for which stability is a common requirement. Perhaps the most important functions of N-glycosylation are to aid in folding of the nascent polypeptide chain and in stabilization of the conformation of the mature glycoprotein. As a consequence, it may also affect any, or all, of the functions that depend on conformation. In the absence of glycosylation, some (glyco)proteins aggregate and/or are degraded, and as a result, are not secreted from the cells in which they are synthesized. Other glycoproteins are less influenced and are secreted, but have compromised biological activities, while some appear to be totally unaffected. Recent studies suggest that one way in which N-linked oligosaccharide chains affect protein folding is by mediating the interaction of the newly-synthesized peptide with calnexin, a chaperone with apparent selectivity for N-glycoproteins [10]. It is a non-glycosylated membrane protein of 65.4 kDa molecular weight, with a large, negatively charged carboxyterminal cytoplasmic tail and an external domain containing three internal repeats of conserved hexapeptide sequence. Experiments with two viral membrane glycoproteins (influenza virus hemagglutinin and vesicular stomatitis virus G protein), showed that calnexin binds transiently to the newly synthesized glycoproteins that have partially trimmed, monoglucosylated oligosaccharides; the binding coincides with protein folding and oligomer assembly. The proteins remain bound to calnexin for different periods of time, depending on the rates at which they achieve conformational maturation. During this time, the single terminal glucose is rapidly turned over in a deglucosylation-reglucosylation cycle. Once a glycoprotein is folded or

Table 1 Carbohydrates modify the properties of the proteins to which they are linked OFTEN Affect solubility, charge and viscosity; Control folding and subunit assembly; Stabilize protein conformation; Protect against proteolysis Affect the lifetime in circulation Change the immunological properties Modify the transmission of signals by cellular receptors Modify the activity of enzymes and hormones RARELY

assembled into oligomeric form, it is no longer reglucosylated and therefore released by calnexin and is free to leave the endoplasmic reticulum. The carbohydrate may change markedly the quartenary structure of a protein to which it is attached, as demonstrated in the X-ray crystallographic study of the Erythrina corallodendron lectin, a member of the large family of legume lectins [ 11]. The heptasaccharide, linked at Asn17 of each of the two subunits of this lectin, prevents the formation of the characteristic dimer observed in other members of the legume lectin family (e.g. concanavalin A and pea lectin). As a result, these subunits adopt a completely different quaternary structure. Not only is glycosylation at a particular site important in directing protein folding and assembly, but the precise structure of the glycan may also be critical. A clear case is that of human chorionic gonadotropin, a glycoprotein hormone composed of two subunits, for which it was shown that abnormally glycosylated a subunit is unable to associate with the [3-subunit to form the mature hormone.

4. MODULATION OF BIOLOGICAL ACTIVITY The ability of carbohydrates to modulate the activities of biologically functional proteins, occasionally even in an all-or-nothing manner, has been established unequivocally during the last decade in a limited number of cases. For most glycoproteins, however, the role of the carbohydrates is still obscure.

4.1. Enzymes Well documented cases on the effect of carbohydrate on enzymatic activity are rare. One of these is that of tissue plasminogen activator (tPA), a serine protease which converts plasminogen into plasmin and thereby induces clot lysis (fibrinolysis). Naturally occurring type I and type II tPA that possess different numbers of N-linked glycans, differ in the rate of formation of an active complex with fibrin that is able to cleave plasminogen. Plasminogen itself exists in two forms, with either one or more oligosaccharide units attached. The rate of fibrin-dependent plasminogen activation spans a range that is dependent on the glycosylation site occupancy of both tPA and its substrate. At the extremes, this activity for type II tPA with

type II plasminogen (possessing one O-linked glycan) is 2-3 times the value for type I tPA and type I plasminogen with one O-linked and one N-linked glycan [ 12, 13]. Another, more recent, case is that of bovine pancreatic RNase. The enzyme occurs both in unglycosylated (RNase A) and glycosylated (RNase B) form; the latter is a collection isoforms, in which the same polypeptide is associated with nine different oligomannose chains (from Man 5 to Man 9) at the single N-glycosylation site (Ash34). RNase A and RNase B have always been reported as having the same enzymatic activity and were frequently quoted as a proof, provided by nature, that such activity is not affected by the presence of carbohydrates in the molecule. With the aid of a novel sensitive assay using double stranded RNA substrate it was shown [13] that RNase A was more than three times as active as RNase B. The individual glycoforms RNase-Mans, RNase-Man 1 and RNase-Man0, prepared by exoglycosidase treatment of naturally occurring RNase B and separated by capillary electrophoresis, were intermediate in activity (Table 2). These differences were attributed to an overall increase in dynamic stability of the molecule with glycosylation (as demonstrated by measurments of proton exchange rates of the various RNase forms) and to steric effects. Molecular modeling indicated that increasing the size of the RNase oligosaccharide up to five mannose residues could lead to a decrease in activity, whereas the Man5 to Man9 glycoforms would exhibit similar activities, as was indeed found [ 14].

Table 2 Enzymatic activity of RNase A and glycoforms of RNase B 1 Relative RNase Carbohydrate activity A None 1.0 B

2

GlcNAc2Man 0 0.62 GlcNAc2Man 1 0.45 GlcNAc2Man 5 0.28 GlcNAc2Mans_9 0.26

t!

1

Based on ref.[ 13].

2

Modified by enzymatic removal of some mannose residues

4.2. Hormones Chemically or enzymatically deglycosylated glycoprotein hormones bind to their receptors on target cells with the same affinity as the native ones; their ability to activate the hormone responsive adenylate cyclase is, however, drastically decreased [reviewed in 2,3]. Site directed mutagenesis experiments on human chorionic gonadotropin cDNA implied that glycosylation at Ash-52 of the (x-chain alone is sufficient for normal signal transduction. Furthermore, in the absence of this critical oligosaccharide unit, glycosylation at Ash-13 of the [3-chain resulted in intermediate activity of the hormone, whereas glycosylation at Ash-30 of the same chain resulted in an inactive product. Deglycosylated hCG interacts with a different domain of the receptor than the native hormone. This difference may be a factor determining the success or failure of signal transduction from the receptor to the effector system.

The role of carbohydrates in the activity of erythropoietin, a glycoprotein hormone that stimulates erythropoiesis, is the subject of intense studies, not the least because of the great commercial interest in this compound. Erythropoietin has the distinction of being the first recombinant glycoprotein produced industrially for clinical use and is being widely employed for the treatment of anaemia in patients on haemodialysis. Desialylation of the hormone enhanced its in vitro activity by increasing its affinity for the receptor, but decreased its activity in vivo, presumably by decreasing its life-time in circulation. Similar results were reported for the N-deglycosylated hormone, which in vitro exhibited several-fold higher specific activity than the native one, but was inactive in vivo. Examination of several preparations of recombinant erythropoietin that differ in the degree of branching of their N-glycans revealed that in vivo activity of the hormone increased with the ratio of tetraantennary to biantennary saccharides [ 15].

4.3. Other biologically active molecules Carbohydrates on receptors may affect the functional coupling of the latter to effector systems such as adenylate cyclase (via guanine nucleotide binding proteins (G-proteins)) and tyrosine kinase, essential for the transmission of signals from the ligand to the cell. Thus, insulin receptor in which all four potential N-glycosylation sites of the b-subunit have been eliminated by site-directed mutagenesis had similar affinity for its ligand as the wild type receptor but lost its transmembrane signaling ability, as evidenced by lack of stimulation of glucose transport and glycogen synthesis by the hormone [ 16]. Two types of T-cell derived, IgE-binding factor have been described, one of which enhances, and the other suppresses, IgE synthesis in mast cells. The factors share a common polypeptide backbone, but only the former is glycosylated. This is perhaps the only known case of such a remarkable change in the activity of a protein caused by the presence of a carbohydrate [ 17]. In IgG antibodies, elimination of the conserved glycan linked to Asn247 of the heavy chain leads to a loss of some of the effector functions of the molecule, such as binding to Fc receptors on macrophages [ 18].

5. ACTIVITIES OF FREE OLIGOSACCHARIDES Diverse activities are exhibited by free oligosaccharides, either derived from glycoproteins or from other sources (Table 3).This was originally demonstrated with heparin oligosaccharides that, like the parent molecule, act as anticoagulants. Recently it was found that a heparin/heparan sulfate dodecasaccharide activates cell-bound fibroblast growth factor, similarly to full-size heparin (or heparan sulfate proteoglycans, such as syndecan) [ 19,20]. The saccharide binds both to the growth factor and its receptor and the formation of such trimeric complex appears to be a prerequisite for signal transduction. Glycans isolated from plant glycoproteins were shown to delay tomato ripening. Other oligosaccharides that act on plants in different ways have been described, some of which are listed in Table 3.

Table 3 Biologically active oli~osaccharides Structure In animals Heparin-derived Heparan sulfate-derived In plants Oligoglucosides Pectin-derived Oligo-GlcN derivatives Fuc-Xyl-N-glycans Xyloglucan-derived

Activity

Ref.'s

Anticoagulant Growth factor activators

[ 19,20] [19,201

Induce disease resistance Anti-auxins Nodulation factors in rhizobia Delay tomato ripening Inhibit or promote elongation of pea stem segments

[21] [21] [22] [231

[21]

6. CARBOHYDRATES AS RECOGNITION DETERMINANTS There is increasing evidence for the concept, formulated over 20 years ago, that carbohydrates act as recognition determinants in a variety of physiological and pathological processes [24,25] (Tables 4 and 5).

Table 4 Carbohydrates and lectins in cell-cell recognition Process Sugars on Infection Host cells Defense Phagocytes Microorganisms Fertilization Eggs Leukocyte traffic Leukocytes Endothelial cells Metastasis Target organs Malignant cells

Lectins on Microorganisms Microorganisms Phagocytes 1 (Sperm) Endothelial cells Lymphocytes Malignant cells (Target organs)

1

Presumed, no experimental evidence available

This concept evolved with the realization that carbohydrates have an enormous potential for encoding biological information. The messages encoded in the structures of complex carbohydrates are deciphered through interactions with complementary sites on carbohydratebinding proteins, chiefly lectins. Processes in which the participation of carbohydrate-lectin interactions was clearly demonstrated include intracellular trafficking of enzymes, clearance of glycoproteins from the circulatory system and a wide range of cell-cell interactions. Particularly exciting is the recent demonstration that binding of carbohydrates on the surface of

leukocytes, with a class of animal lectins designated selectins, controls leukocyte traffic by mediating adhesion of these cells to restricted portions of the endothelium and their recruitment to inflammatory sites. 6.1. Clearance (traffic) markers The rapid removal of desialylated glycoproteins from rabbit serum via the hepatic asialoglycoprotein receptor (or lectin), a phenomenon discovered in the late 1960's, is the prototype of the saccharide-based recognition system, although its role in nature has not yet been proven beyond doubt [26].

Table 5 Clearance and targeting of glycoproteins Glycoprotein Specificity Asialoglycoproteins Galactose Hormones SQ-GalNAc Lysosomal enzymes Diverse

Receptor

Man-6-phosphate Mannose

Location Liver (hepatocytes) Liver (Kupffer cells, endothelial cells) Ubiquitous Macrophages, liver(endothelial cells)

Several other systems in which the traffic of glycoproteins is controlled by their carbohydrate constituents are known (Table 5). A prominent example is the intracellular routing of lysosomal enzymes to their compartment which is mediated by the recognition between Man-6-P attached to the oligomannose unit(s) of such enzymes, and the Man-6-P receptors [4]. Two such receptors have been described, one cation-independent and of high molecular weight (220 kDa), the other cation-dependent and of low molecular weight (48 kDa). A defect in the synthesis of the Man-6-P marker recognized by the receptors results in I-cell disease (also called mucolipidosis II or MLII), an inherited lysosomal storage disease, characterized by a lack in the lysosomes of those enzymes that normally carry the marker [27]. It is caused by a deficiency of GlcNAc-phosphotransferase, the first enzyme in the pathway of mannose phosphorylation, and is thus a processing disease, the first of its kind to be identified. Therefore, even though the disease is transmitted by a single gene, some 20 enzymes are affected. The enzymes lacking the recognition marker do not reach their destination (the lysosomes), and are, consequently, secreted into the extracellular milieu, which is one of the biochemical abnormalities of the affected cells. The specificity of the GlcNAcphosphotransferase for certain lysosomal enzymes is based on its ability to recognize a specific lysine residue and a particular tertiary domain of the acceptor glycoprotein. Another carbohydrate-specified targeting system is that of the sulfated glycoprotein hormones. Native lutotropin (LH), carrying predominantly mono- and di-sulfated oligosaccharides on its [3-subunit, is cleared from the circulation 4-5 times more rapidly than recombinant LH, produced in CHO cells, that bears only sialylated oligosaccharides. The sulfated oligosaccharides of LH are synthesized by the action of two enzymes, a glycoprotein

hormone-specific GalNAc-transferase and a GalNAc-specific sulfotransferase [28]. The oligosaccharide formed by the two transferases, SO4-GalNAc~4GlcNAc132Manot is recognized by a receptor present on hepatic endothelial and Kupffer cells. Sulfated oligosaccharides are also present on the common precursor to two other hormones, adrenocorticotropin and melanotropin. It has been hypothesized that the attachment of this structure is a general tag that signals rapid clearance, resulting in a short burst of circulating hormones, thus preventing overloading of the corresponding receptors. The presence of well-defined carbohydrate binding proteins on cell surfaces is being exploited for drug targeting to specific organs. Gaucher's disease is caused by a deficiency of the enzyme 13-glucocerebrosidase, resulting in accumulation of glucocerebroside in Kupffer and endothelial (non-parenchymal) cells of the liver. These cells contain on their surface a mannose- (and N-acetylglucosamine) specific lectin. To target the 13-glucocerebrosidase into the above cells, the complex and hybrid sugar chains of the enzyme were trimmed down with the aid of exoglycosidases to expose the mannose residues of the pentasaccharide core. In this way the administered glucocerebrosidase is effectively delivered to the deficient cells where the enzyme is needed to degrade the accumulated glucocerobroside [29]. 6.2. Infection The oligosaccharide repertoire on the host-cell surface is among the key genetic susceptibility factors in viral and microbial infection and in toxin action. A number of viral, mycoplasmal, bacterial and protozoan pathogens use specific carbohydrate structures (of glycoproteins or glycolipids) on host cells as attachment sites in the initial stages of infection [30,31]. Experiments in intact animals have indeed proved that it is possible to prevent bacterial infection by blocking the attachment of the responsible organism with an appropriate sugar (Table 6). Such findings have provided an impetus for the development of carbohydratebased anti-adhesion drugs to combat infections. At present, at least two drugs of this kind have been patented, against bacterial pneumonia and Helicobacter pylori, (a bacterium associated with stomach ulcers). Should a bacterium mutate so that it no longer recognizes the antiadhesive carbohydrates, it will also fail to bind to its cell surface receptors, and therefore lose the ability to cause infection. Moreover, since such drugs do not kill the pathogens, they will not exert selection pressure and their use will not result in the development of resistance. Even if a particular carbohydrate has been established as an inhibitor for a disease-causing microorganism in an animal (or in humans), it must be determined whether the use of this carbohydrate, or its analogues, will not interfere with other processes in the body. One such process is lectinophagocytosis, well documented for the mannose specific E. coli [32]. This mode of phagocytosis may result from binding of the bacteria to phagocytes, e.g. macrophages or neutrophils, which is followed by activation of the phagocytes and uptake and killing of the bacteria. Lectinophagocytosis may occur in vivo and may provide protection against infection by bacteria to nonimmune hosts or in sites that are poor in opsonins. The latter include lungs, renal medulla, the cerebrospinal fluid and the peritoneal cavity, especially during peritoneal dialysis. In another mode of lectinophagocytosis, a wide range of microorganisms (bacteria, fungi and protozoa) that express mannose on their surface, bind to the mannose specific lectin present on the surface of macrophages. This binding, too, may lead to the uptake of the

10 microorganisms by the phagocytic cell and occasionally also their killing. A particularly interesting example of such a microorganism is the pathogenic fungus, Pneumocystis carinii, a

Table 6 Inhibitors of sugar-specific adhesion prevent infection in vivo Organism Animal, site Escherichia coli type 1 Mice, UT Mice, GIT Mice, UT Klebsiella pneumoniae type 1 Rats, UT Shigella flexnerii type 1 Guinea pigs, eye Escherichia coli type P Mice Monkeys Escherichia coli K99 Calves. GIT Pseudomonas aeruginosa UT, urinary tract; GIT, gastrointestinal tract

Human, ear

Inhibitor Meo~Man Mannose Anti-Man antibody Meo~Man Mannose Globotetraose GalI34Gal13OM Glycopeptides of serum glycoproteins Gal+Man+NeuAc

major case of death among AIDS patients. Human immunodeficiency virus (HIV), the causative agent of AIDS, is heavily glycosylated [reviewed in 2]. The major envelope glycoprotein gpl20 of the HIV has a key role in infection by the virus through its interaction with the membrane glycoprotein CD4 of T lymphocytes. Glycosylation of gpl20 appears to be a prerequisite for CD4 binding: the non-glycosylated protein from cells grown in the presence of tunicamycin does not bind to CD4 and treatment of gpl20 with deglycosylating enzymes impairs binding. Of the various inhibitors of glycosylation tested, the most dramatic anti-viral effects observed have been with N-butyldeoxynojirimycin.

6.3. Leukocyte traffic Research carried out mainly during the last 5 years has demonstrated that adhesive interactions mediated by surface carbohydrates and surface lectins play a crucial role in leukocyte trafficking to sites of inflammation and hemostasis and in the migration (homing) of lymphocytes to specific lymphoid organs. In these processes, the carbohydrates serve as "area codes" which are interpreted by E-selectin, P-selectin and L-selectin, members of a family of endogenous lectins [33-35]. The selectins are highly asymmetric membrane-bound proteins. Their extracellular part consists of an amino terminal carbohydrate recognition domain (CDR), an epidermal growth factor-like domain and of several short repeating units related to complement-binding protein They bind specifically to sialyl-Lewisx (siaLe x in brief), NeuAc(ot2-3)Gal(131-4)[Fuc(o~l-3)]GlcNAc, and its positional isomer, sialyl-Lewis a (siaLea), NeuAc(o~2-3)Gal(131-3)[Fuc(o~l-4)]GlcNAc, with both fucose and sialic acid required for binding; sialic acid can be replaced by another negatively charged group such as sulfate. These proteins recognize the carbohydrate ligands only when the latter are present on particular

11 glycoproteins, such as cell surface mucins, pointing to the role of the carrier molecule and carbohydrate presentation in the recognition of the latter by lectins. The selectins provide the best paradigm for the role of sugar-lectin interactions in biological recognition. In broad outline, they all mediate, although with some differences, the adhesion of circulating leukocytes to endothelial cells of blood vessels, leading to the exit of the former cells from the circulation. The extravasation is necessary for the migration of leukocytes into tissues, such as occurs under normal recirculation of lymphocytes between different lymphoid organs or in recruitment of leukocytes to sites of inflammation. L-selectin, also known as "homing receptor", is found on all leukocytes. It is predominantly involved in the recirculation of lymphocytes, directing them specifically to peripheral lymph nodes. In contrast to the homing receptor the two other selectins are expressed mainly on endothelial cells, and only when these cells are activated by inflammatory mediators, mainly cytokines (e.g. interleukin-2 and tumor necrosis factor). The latter are released from tissue leukocytes in response to e.g wounding, infection or ischemia and induce the expression of P-selectin on the endothelial surface within minutes and of E-selectin within 3-4 hours. Recent experiments in animals have provided direct evidence for the role of selectins in the control of leukocyte traffic. For instance, in P-selectin deficient mice, generated by targeted gene disruption, the recruitment of neutrophils to the inflamed peritoneal cavity was significantly delayed [36]. The clinical importance of selectin-carbohydrate interactions in acute inflammatory responses in humans is illustrated by the finding that the neutrophils of two patients had a deficiency in SiaLe x [37]. The specific biochemical lesion responsible for this defect has not yet been established, but is believed to be a reflection of a general fucosyltransferase deficiency in these patients. In agreement with the ligand activity of SiaLe x, the neutrophils of the patients were unable to bind to E-selectin. The above findings imply that this inability prevents the neutrophils from migrating to the sites of infection and suggest that inhibitors of the selectins may be potent anti-inflammatory agents. Prevention of adverse inflammatory reactions by inhibition of leukocyte extravasation has become a major aim of many pharmacological industries. Preliminary experiments in animal models indeed show that oligosaccharides recognized by the selectins exert protective effects against experimentally induced lung injury. These approaches are now being evaluated for treatment of human disease.

7. OPEN QUESTIONS Although the enormous progress made in the last decade in many aspects of glycobiology "has finally opened a crack in the door to one of the last great frontiers of biochemistry" [38], many questions remain unanswered. The structures of the glycans encountered are exceedingly diverse and we are, as yet, unable to discern the principles that guide their formation. Although preliminary insights have been obtained into the central role of the polypeptide backbone in specifying glycosylation [28], the mechanism(s) by which the information encoded in the primary amino acid sequence is translated into a particular glycan structure is still not clear. A related problem concerns the molecular basis for the variations in glycosylation of the same protein between different species or cell types, and in the course of development, differentiation and oncogenesis.

12 The intriguing question of the function(s) of the carbohydrate is for most glycoproteins, whether soluble or membrane bound, still wide open, since only for a small number of glycoproteins, modulation of physicochemical properties or biological activities by their glycans has been demonstrated. One thing is clear: glycosylation can have markedly different effects on different proteins. This means that each glycoprotein must be examined individually and meticulously for the possible functions of the glycans it carries. Progress in this area will therefore be unavoidably slow. Still unresolved is the question of the biological relevance of microheterogeneity - in other words, can unique biological functions be ascribed to different components of the ensemble of glycoforms? The accumulation of evidence on the role of carbohydrates, whether bound to proteins or lipids, as recognition molecules is exceedingly rewarding. It serves as a strong impetus for improving techniques of structural analysis of carbohydrates as well as for the development of new methods of synthesis of oligosaccharides and glycomimetics, a challenge for the carbohydrate bioengineers. Recent developments also focus attention on lectins as co-partners in the recognition process. Of particular importance in this context is the need for a detailed knowledge of their combining sites [39], which should allow the design of highly effective carbohydrate anti-adhesive drugs.

8. A C K N O W L E D G E M E N T Special thanks are due to Dr. Halina Lis for her help in preparing this manuscript.

9. REFERENCES

1

N. Sharon, Complex Carbohydrates: Their Chemistry, Biosynthesis and Functions. Addison-Wesley, Reading, Massachussets, 1975. 2 H. Lis and N. Sharon, Eur. J. Biochem., 218 (1993) 1. 3 A. Varki, Glycobiology, 3 (1993) 97. 4 S. Kornfeld, Annu. rev. Biochem., 61 (1992) 307. 5 M.N. Fukuda, Glycobiology, 1 (1990) 9. 6 J. Jaeken, H. Carchon and H. Stibler, Glycobiology, 3 (1993) 423. 6a J. Jaeken, H. Schachter, H. Carchon, P. De Cock, B. Coddeville and G. Spik, Arch.Dis. Child., 71 (1994) 123. 7 B.D. Shur, Curr. Biol., 4 (1994) 996. 8 E. Ioffe and P. Stanley, Proc. Natl. Acad. Sci., USA 91 (1994) 728. 9 M. Metzler, A. Gertz, M. Sarkar, H. Schachter, J.W. Schrader and J.D. Marth, EMBO J., 13 (1994) 2056. C. Hammond and A. Helenius, Curr. Biol., 3 (1993) 884. 10 11 B. Shaanan, H. Lis and N. Sharon, Science, 254 (1991) 862. 12 A.J. Wittwer, S.C. Howard, L.S. Carr, N.K. Harakas, J. Feder, R.B. Parekh, P.M. Rudd, R.A. Dwek and T.W. Rademacher, Biochemistry, 28 (1989) 7662. 13 P.M. Rudd, H.C. Joao, E, Coghill, P. Fiten, M.R. Saunders, G. Opdenakker and R.A. Dwek, Biochemistry, 33 (1994) 17.

13 14 15 16 17 18 19 20 21 22 23 24 25 26

27 28 29 30 31 32 33 34 35 36 37 38 39

R.J. Woods, C.J. Edge and R.A. Dwek, Nature Struct. Biol., 1 (1994) 499. M. Takeuchi and A. Kobata, Glycobiology, 1 (1991) 337. I. Leconte, C. Auzan, A. Debant, B. Rossi and E. Clauser, J. Biol. Chem.,267 (1992) 17415. K. Ishizaka, Annu. Rev. Immunol., 6 (1988) 513. R.A. Dwek, Glycoconjugate J., 10 (1993) 357. D. Spillman and U. Lindahl, Curr. Opin. Struct. Biol., 4 (1994) 667. A.D. Lander, Chemistry&Biology, 1 (1994) 73. A. Darvill, C. Augur, C. Bergman et al., Glycobiology, 2 (1992) 181. I. Vijn, L. das Neves, A. van Kammen, H. Franssen and T. Bisseling, Science, 260 (1993) 1764. H. Yunovitz and K.C. Gross., Physiologia Plantarum, 90 (1993) 152. N. Sharon and H. Lis, Sci. Am., 268(1) (1993) 82. T. Feizi, Curr. Opin. Struct. Biol., 3 (1993) 701. G. Ashwell, in Lectin Blocking: New Strategies for the Prevention and Therapy of Tumor Metastasis and Infectious Diseases (J. Beuth and G. P Pulverer, eds.), Gustav Fischer Verlag, Stuttgart Jena New York, 1994, p. 26. E.F. Neufeld, Annu. Rev. Biochem., 60 (1991) 257. J.U. Baenziger, FASEB J., 8 (1994) 1019. E. Beutler, A. Kay, P. Garver, D. Thurston, A. Dawson and B. Rosenbloom, Blood, 78 (1991) 1183. I. Ofek and N. Sharon, Curt. Top. Microbiol. Immunol., 151 (1990) 91. I. Ofek and R.J. Doyle, Bacterial Adhesion to Cells and Tissues. Chapman and Hall New York London 1994 I. Ofek, J. Goldhar, Y. Keisari and N. Sharon, Annu. Rev. Microbiol., 49 (1995) 239. L.A. Lasky, Science, 258 (1992) 964. M.P. Bevilacqua, Annu. Rev. Immunol., 11 (1993) 767. S.D. Rosen and C.R. Bertozzi, Curr. Opin. Cell Biol., 6 (1994) 663. T.N. Mayadas, R.C. Johnson, H. Rayburn, R.O. Hynes and D.D. Wagner, Cell, 74 (1993) 541. U.H. von Andrian, E.M. Berger, L. Ramezani, J.D. Chambers, H.D. Ochs, J.M. Harlan, J.C. Paulson, A. Etzioni and K.E. Arfors, J. Clin. Invest., 91 (1993) 2893. G.W. Hart, Curr. Opin. Cell Biol., 4 (1992) 1017. N. Sharon, Trends Biochem. Sci., 18 (1993) 221.

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S.B. Petersen, B. Svensson, and S. Pedersen (Eds), CarbohydrateBioengineering 9 Elsevier Science B.V. All rights reserved.

15

NMR studies of the structure and dynamics of carbohydrates in aqueous solution Herman van Halbeek and Shuqun Sheng Complex Carbohydrate Research Center and Departments of Chemistry and Biochemistry, The University of Georgia, 220 Riverbend Road, Athens, Georgia 30602-4712, USA

Abstract

Notable recent developments in NMR methodology for studying carbohydrate structure and dynamics include the increased information gained from the observation of hydroxyl proton signals in supercooled water as well as improved measurements of long-range heteronuclear scalar couplings and 13C relaxation rates. This chapter presents an example of recent progress in each of those categories. (1) A 2-D homonuclear rotating-frame exchange experiment is proposed as a suitable means under supercooled aqueous conditions to detect transient intramolecular hydrogen bonds in the flexible disaccharide sucrose. (2) A modified 2-D 1H-detected heteronuclear multiple-quantum experiment is introduced for speedy and accurate measurement of nJcH (n>2); the application of the so-called HSMBC experiment is illustrated for sialyllactose. (3) The quantitative aspects of the dynamics of the hexasaccharide headgroup of ganglioside GDla embedded in a perdeuterated mixed micelle in aqueous solution are assessed by 1H-detected natural-abundance 13C Tlo measurements; the data are interpreted according to the Lipari-Szabo model-free approach which reveals relatively fast internal motions in the outer region of the hexasaccharide.

1. INTRODUCTION A detailed NMR analysis of the solution conformations and dynamics of a carbohydrate encompasses the following steps: 9 complete assignment of 1H and 13C NMR spectra of the carbohydrate; 9 establishment of spatial (distance and/or torsion angle) constraints between atoms from dipolar and scalar correlation NMR measurements; 9 measurement of 1H and 13C relaxation parameters (that is, T1, T2, Tip, homo- and heteronuclear cross-relaxation rates). Despite the efforts typically involved in conducting the pertinent NMR experiments [ 1,2], the experimentally measurable constraints may fall short in determining the complete conformation and dynamic behavior of an oligosaccharide. Evaluation of the experimental data with computational strategies is then a necessity, typically by using potential energy calculations and molecular dynamics (MD) simulations [3-5]. However, in favorable cases, the NMR study yields a sufficiently large number of constraints to broadly define the conformation of the oligosaccharide in aqueous solution. Often one finds that not all the

16 obtained NMR constraints are compatible with the existence of a single rigid structure, implying that the oligosaccharide is dynamic with respect to torsional vibrations around each glycosidic bond. The past few years have witnessed a vast increase in the number of efforts aimed at the measurement of NMR parameters directly related to the flexibility of carbohydrates, including 1H and 13C T1 and Tip, homo- and heteronuclear cross-relaxation rates, and global and local correlation times (reviewed in [6]). It is gradually becoming clear that very few oligosaccharides adopt a single, fully constrained ("rigid") conformation. The ability of most carbohydrates to present one covalent structure to their environment in many different ways may very well contribute to their versatility in biological functions. NMR spectroscopy not only helps to narrow the conformational space accessible to a flexible oligosaccharide, it also can provide information about the relative populations of and the rate of interconversion between different energetically favored conformers. We will discuss below three examples of this role for NMR in carbohydrate conformational analysis, namely, to aid in restricting the theoretically possible ensemble of conformations for a given oligosaccharide. We present a couple of recently developed NMR methods that provide either additional (hydrogen bond) or more accurately determined (3JcH) parameters to restrain the oligosaccharide structure in question. Also, a quantitative evaluation of 13C relaxation data is presented for a glycolipid system mimicking physiologic cell-surface conditions.

2. HYDROGEN BONDING Hydroxyl proton resonances in aqueous solutions of sugars were first observed over 15 years ago [7], but the value of these protons in the conformational analysis of carbohydrates has been demonstrated only recently [8-11]. Hydroxyl resonances have the potential to provide a wealth of structural information in the form of chemical shifts, 3JH_c_OH couplings, nuclear Overhauser effects (NOEs), and exchange rates. However, this information is accessible only if the intermolecular exchange of OH protons with solvent H20 can be slowed sufficiently. At room temperature, hydroxyl protons in aqueous solutions of carbohydrates engage in chemical exchange with water, the rate of which is very fast on the NMR time scale. Researchers at first applied mixed solvents (water/acetone and water/methanol) to study OH groups at low temperatures (-5 to -10~ (see, e.g., [9]). More recently, 1H NMR studies have been reported of hydroxyl groups in dilute solutions of monoand disaccharides in pure H20 under supercooled (-15 t o - 2 0 ~ conditions [10]. The chemical exchange rates under these conditions are reduced to such an extent that signals can be observed for each hydroxyl site; thus, all hydroxyl proton resonances can be assigned on the basis of (scalar or dipolar) connectivities to non-labile aliphatic protons. The line widths, temperature shift coefficients, and coupling constants of OH protons are valuable hydration and hydrogen-bonding probes in NMR studies [11]. Also, H/D isotope effects of hydroxyl protons/deuterons on 13C resonances can be used to obtain indirect evidence of the involvement of OH groups in intra- or interresidue hydrogen bonds [12]. Furthermore, protruding farther from the glycosyl ring systems than most CH protons, OH protons may serve as long-range sensor conformational probes which can be interrogated by NOESY and ROESY experiments on the carbohydrate in aqueous solution [8,9]. In order to use OH protons as conformational probes for oligosaccharides, special NMR techniques for water

17 suppression must be utilized [13-16]. We report below on the study of intramolecular hydrogen bonds in the disaccharide sucrose in aqueous solution. The three-dimensional structure of sucrose [Fruf-13(2~-~1)~-Glup] (Fig. 1), particularly the conformation of its glycosidic linkage, has been the subject of numerous investigations. Data obtained by NMR analyses and MD simulations on sucrose in solution have led to uncertainties concerning the degree of rigidity of the linkage conformation in solution [ 17-21 ], raising the question whether it is the same as in the crystal structure [22]. Careful 13C T1 measurements for sucrose revealed very fast and small-amplitude torsional and vibrational motions at different sucrose ring positions [23,24] and mobilities of the exocyclic groups different from the ring skeletons [25]; however, on the basis of this type of measurements the glycosidic linkage was judged to be rigid and similar in conformation to that observed in the crystalline state. In 1992, inspired by the work of P6rez c . s . [20], we reinvestigated the solution conformation of sucrose applying new NMR methods to reveal 1H/1H NOE contacts explicitly including OH protons [26]. The results of our quantitative NOE measurements are compiled in Table 1. Interestingly, all NOE connectivities found for sucrose in water solution can be explained in terms of a single conformation, which is virtually identical to the crystal structure. However, based on a quantitative analysis of the magnetic field strength dependence of 1H/1H NOE data, we questioned [26] the rigidity in solution of the sucrose glycosidic bond. The NOEs between protons on different rings proved to be magnetic field strength-dependent, while this did not seem to be the case for NOEs between protons within the glucopyranose ring. We interpreted this observation as evidence of rearrangements occurring around the glycosidic bond between the glucosyl and fructosyl residue that take place much more quickly than the tumbling rate of the molecule in solution. This type of internal motion is transparent to relaxation parameters (including 13C T1) if it occurs on the same time scale as overall molecular tumbling; this seems to be the case for sucrose at ambient temperature. Thus, we demonstrated that sucrose in solution more than likely experiences fast internal motion around its glycosidic bond.

Figure 1. Left: the crystal structure of sucrose (hydrogen bonds are represented by dashed lines) [22]. Right: the 600-MHz 1-D 1H NMR spectrum of sucrose in pure water, recorded at -17~ and pH 6.5.

18 Table 1. Apparent transglycosidic interproton distances for sucrose in aqueous solution Proton pair a Hlg/Hlf Hlg/H3f Hlg/H4f Hlg/H6f H5g / H4f Hlg / OHlf H 1g / OH6f H l g / OH3f H5g / OH3f OH2g / H 1f

Distance b (,&) 2.6 4.6 3.5 4.0 3.0 3.6 3.4 4.0 3.8 3.2

a Data were derived from NOE contacts

observed at 500 MHz and 27~ for CH, and at-10~ for OH protons [26]; 'g' denotes the glucosyl, 'f' the fructosyl ring. b Distances were calculated using the formula rij=((Yref/(Yij)l/6rref in which (Yrefand rre f represent the cross-relaxation rate of and the distance between a reference pair of protons. Intraglucosyl distances H lg/H2g (2.4 ]k), Hlg/H3g (3.7 A) and Hlg/H4g (4.0 ]k) were used as references.

Table 2. Temperature coefficients and scalar coupling constants of the hydroxyl protons of sucrose dissolved in pure water 3JH_c_OH c

Kb

Proton a

OHlf OH3f OH4f OH6f OH2g OH3g OH4g

-8.2 -8.8 -9.2 -11.0 -8.0 -10.3 -9.4

5.7 7.3 5.6 5.3 7.4 4.8 6.4

OH6g

-10.3

5.5

a ,f, denotes the fructosyl, 'g' the glucosyl ring. b Temperature coefficients (~) are in p p m / ~ 103; they were measured over the temperature range-18 to-8~ at 1~ intervals. c Scalar coupling constants (3Jn_c_on) are in Hz; the values were measured a t - 17~ OH H Ha\ ..Hb

3'1

H

-OH '

t.,

H ':2

"

..HOI ~ 0 1C H2

5 C H20H 6

In this study we refine our investigations of sucrose at low temperatures in aqueous solutions in an effort to shed light on the existence of any interresidue hydrogen bonds in the context of its newly revealed flexibility around the glycosidic bond. We were aware that a great deal of effort had been spent by others [27,28] examining intramolecular hydrogen bonding and determining its effect on the overall conformation of sucrose in solution. However, we are the first to use the hydroxyl proton signals of sucrose in supercooled aqueous solution as NMR probes for the detection of intramolecular hydrogen bonds. A capillary tube (1.5 mm i.d.) containing a 20 mM sucrose solution in HzO/D20 (9:1, v/v) was inserted into a 5-mm tube and, once placed in the magnet, the sample was gradually cooled to -17~ All of the sucrose OH signals were indeed observed in 1-1 echo water suppression [ 13] experiments (Fig. 1) and had lines sufficiently narrow to be assigned by a 2-D COSY experiment. Neither the OH temperature shift coefficients (see Table 2), 3JH_C_OH scalar couplings (see Table 2), 13C (COH v s . COD) chemical shifts, nor OH exchange rates allowed us to single out any sucrose hydroxyl group with characteristics significantly different from the others. However, 2-D rotating-frame exchange spectroscopy (ROESY) of sucrose under supercooled conditions revealed a direct exchange between the glucosyl OH2 and fructosyl OH1 protons (see Fig. 2). We conclude that the observation of chemical exchange between OH2g and O H l f ('g' denotes the glucosyl, 'f' the fructosyl moiety in sucrose) provides reasonable

19 evidence for the transient existence of an OHlf:::O2g (or OH2g:::Olf) hydrogen bond in solution under the applied supercooled conditions. This "weak" hydrogen bond is compatible with the flexibility of sucrose around its glycosidic bond. It appears that the intramolecular hydrogen bonding in sucrose is concentration dependent; the existence of hydrogen bonds OHlf:::O2g, OH3f:::O2g, and OH6f:::O5g at higher concentrations has been postulated based i.a. on 13C isotope effects [27]. The existence of the first two hydrogen bonds at high concentration in solution implies flexibility and fast interconversion between two conformers around the glycosidic linkage in solution; in the crystal state only OHlf:::O2g and OH6f:::O5g are observed. It should be noted that the aforementioned 1H/1H ROESY experiment can only detect direct chemical exchange between two OH groups, so it is restricted in its applicability to those hydrogen bonds that involve two participating OH groups (thus, ROESY cannot reveal any hydrogen bonding to ring oxygens). Moreover, the ROESY spectrum contains no information about the identity of the hydrogen-bond donor and acceptor and, therefore, cannot discriminate between the possibilities of an OHlf:::O2g and an OH2g:::O 1f hydrogen bond in sucrose.

I

2g/!f@ f

o

E Q_ EL IY)

-~;

4g

Q_ i

(D --

~

(.o

(~4f 6.6 1

6.~ 6.0 F2 (ppm) I

I

Figure 2. Hydroxyl proton region of a 2-D ROESY spectrum of sucrose in H20/D20 (9:1, v/v) recorded at 600 MHz, pH 6.5, and-17~ The effective spin-lock field strength was set to 1.5 kHz, and the ROESY mixing time was 100 ms.

20

3. HETERONUCLEAR LONG-RANGE COUPLINGS MEASURED BY A TWODIMENSIONAL H.S.M.B.C. EXPERIMENT The classic HMBC experiment [29] has long been recognized as a powerful means to qualitatively detect long-range heteronuclear couplings. For carbohydrates, the HMBC experiment provides the key to primary sequencing, exclusively and entirely based on tracing through-bond J-couplings, the only scalar couplings consistently observable across a glycosidic bond being the transglycosidic 3JcH [30]. However, determining from the HMBC cross-peak patterns the magnitudes of the involved nJcHwith sufficiently high accuracy is not straightforward. Cross-peak multiplets in HMBC spectra are recorded in mixed-phase mode in the 1H dimension due to evolution of 1H chemical shifts and homonuclear couplings during the relatively long delay period required for heteronuclear long-range couplings to evolve [31]. Direct extraction of heteronuclear coupling constants from non-pure absorption patterns is impossible. To find a reference spectrum of the oligosaccharide that has the same phase behavior as the cross peak in the HMBC spectrum is mandatory. A 2-D 1H/1H TOCSY spectrum recorded under otherwise the same conditions as the HMBC spectrum has been used successfully for this purpose [32]. Alternatively, the refocused HMBC experiment designed for the quantitative measurement of long-range coupling constant values based on the HMBC cross-peak intensities [33,34] uses either a 1-D version of the HMBC spectrum (obtained by omitting the 13C pulses from the original sequence) or a full 2-D HMQC spectrum as reference. Both approaches require, therefore, the recording of the reference spectrum in an experiment separate from the HMBC experiment itself. The method we propose here uses the original HMBC pulse sequence, without suppression of one-bond correlations. Indeed, the one-bond correlation peaks are essential because they provide the necessary references for the phase properties of the multiple-bond correlations. Hence, we refer to this experiment as HSMBC for heteronuclear single- and multiple-bond correlation spectroscopy. The pulse sequence of the 2-D HSMBC experiment is shown in Fig. 3. No separate, additional NMR experiment needs to be conducted to obtain an appropriate reference spectrum. Also, the reference peaks here have truly the same phase properties as the multiple-bond correlation peaks, as they are collected during the very same experiment. Values for the long-range couplings can be extracted applying the shift method introduced by Titman et al. [32]. We will illustrate the application of the HSMBC experiment for sialyl(z(2~6)-lactose (SL6):

H

oH

H

Ho

Hn ~9~c~H .

" " C . . " K 17 16 H C

ILI

o

H

21,~

OH

g

[

.~

6H

i4)~ ~

H

O

O/,\

oH ~

\

, "

n o - ~

H

Neu5 Ac- c~(2~ 6)- Galp- 13( 1-+4)- Glcp

OH

21

90x

1H

180x

~zx

tl/2

II

90t)l

Figure 3.

tl/2 9002

Pulse sequence of the 2-D HSMBC experiment. The phase cycling is as follows:

~)1 = X,-X; (~2 = X, X, X, X,--X,--X,--X,--X; t~3 = X,--X, X,--X,--X, X,--X, X.

The HSMBC spectrum of SL6 (Fig. 4) was recorded on a Bruker AMX-600 spectrometer (1H frequency 600 MHz, 13C frequency 150 MHz) using a 20 mM solution of the trisaccharide in D20 at pD 7.3 and 25~ The delay A was set to 45 ms, and the acquisition times t 2 and t 1 were 1.13 s (spectral width 1805 Hz, 4 K complex data points) and 25.6 ms (spectral width 10,000 Hz, 512 real data points), respectively. The relaxation delay was 1 s; 64 scans were accumulated per t~ increment. Quadrature detection in t I was accomplished by timeproportional phase incrementation. The total duration of the HSMBC experiment on SL6 was 15 h. The HSMBC data were processed using BioSym's FELIX software package, version 2.3, on a Silicon Graphics Personal Iris workstation. No multiplication by any window function and no zero-filling in t2 were applied before Fourier transformation. A cosine-squared window function was applied in the tl dimension; the first data point was multiplied by 0.5 to reduce t l noise. No zero-filling was applied in t l. The final resolution in the 13C dimension was 19.5 Hz/pt. Fourier transformation in both dimensions was followed only by phasing in F1. The pair of resulting 1H cross sections was selected that contain the 3JcH and 1JcH correlation multiplets for a particular proton. Each of these spectra was then subjected to Hilbert transformation (to create the imaginary part of the spectral slice) followed by inverse Fourier transformation; subsequent zero-filling to 16 K real points resulted in a digital resolution of 0.1 Hz/pt. After Fourier transformation, the baseline of the signals was carefully adjusted applying the "flat" routine in FELIX and any unwanted signals in the traces were zeroed. The selected 1JcH multiplet was then inverted, shifted by 0.5x3J(trial), and added to its parent multiplet which was shifted in the opposite direction by 0.5x3J(trial). The resulting convoluted multiplet was fitted to the experimental 3JcH multiplet. The calculations are performed by an in-house FORTRAN software routine on a DEC-3100 workstation. The best fit is achieved when COS ~ --

E.T

IElITI reaches its maximum [35], where E is the vector corresponding to the experimental 3JcH multiplet and T is the trial vector reconstructed from the reference multiplet by the procedure outlined above.

22

Neu H3e Neu H3a

Glccz H 1Gal H 1

.!

C) ! |

,0.

.o,

-o

i "~'

o t

Neu C5.b w . , q

,,, ,,

Neu C4.Glccz C5 ~ GIc C4~ Glccz C1~ Glcl3 C1-.Neu C2-,Gal C1/-"

o

E Q_

-o

Q-

,,,

,e.,

N

.,,

.-.

,,,~ll0Do.o

D

o,

(.O ~--,

,.

,....

.,

.-,

,D

..

9

9

..

O

9

LL_

-o

00

i

II

cb

,

!

El

-o

}, I

5.0

I

4.0 F2

I

3.0

(ppm)

Q

o

I

2.0

Figure 4. 2-D HSMBC spectrum of sialyl-(z(2-*6)-lactose in D20, recorded at 600 MHz, pD 7.3, and 25~

The procedure for obtaining the value of a particular long-range coupling constant is graphically illustrated for Glc(z 3JH1c5. Figure 5 shows the doublet taken from the one-bond Glccz 1JH1c1 trace of the HSMBC spectrum, and the subsequent reconstruction of the 3JH1c5 multiplet. The 3JH1c5 coupling constant obtained by this method is 6.5 Hz. This value is accurate to _+0.2 Hz, that is, within two times the digital resolution of the 1-D traces processed as described above. Figure 6 shows the matching patterns of five other nJcH multiplets for SL6, including those for the interglycosidic coupling 3JGa1H1-GlcC4- The values obtained by our method are marked in bold; the italicized values are those obtained by a quantitative HMBC experiment [34]. The selected subset of nJcH values for sialyllactose illustrates that the HSMBC method provides generally more accurate values for the desired couplings in a shorter period of time. The need to record a separate reference spectrum is circumvented, while the accuracy of the "Jfn values is significantly improved over the quantitative HMBC experiment. The HSMBC method also compares favorably with 1-D selective HSQC measurements of the same couplings in SL6 [36].

23 (a)

(b) (c)

31 I~I~, _1

'atrial /~~ [ 'aH1-H2

/v L_

(d) Figure 5. The J-fitting procedure as performed for Glco~ 3JH1c5 from the pertinent traces in the 2-D HSMBC spectrum of sialyl-~(2~6)-lactose. (a) Doublet of Glc~ H1 taken from the 1JH1c1 row; (b) inverted doublet 'a' left-shifted by 3Jnlcs(trial); (c) reconstructed multiplet obtained by co-adding 'a' and 'b'; (d) actual multiplet of Glc~ H1 taken from the 3JHlc5 row.

Gal H1-GIc C4

4.0

(3.9+_0.8)

Neu H3a-C5

+i

Neu H7-C8

Neu H3a-C4 (7.7+ 1.1)

Neu H3a-C2

io'

. ._+ . ~f~

Glccz H1-C5 65

('4.0__+0.+4)

(6.5+_0.9)~

Figure 6. Selected set of 2JcH and 3JcH couplings (__.0.2 Hz) for sialyl-a~(2~6)-lactose, resulting from the fit of experimental (bottom traces) and reconstructed (top traces) 2-D HSMBC cross-peak multiplets. The resulting coupling constant values (in Hz) are marked in bold. For comparison, the values determined by Zhu et al. [34] from a quantitative HMBC experiment on SL6 are included (in italics).

24 4. DYNAMICS OF MEMBRANE-BOUND CARBOHYDRATES

The molecular behavior of an oligosaccharide chain isolated from a glycoprotein or glycolipid hardly mimics the behavior of the chain when bound to a protein or a lipid. For example, the molecular motion of a freely tumbling oligosaccharide will be substantially different from that of the same oligosaccharide anchored to a protein backbone and/or in a lipid bilayer. It is likely that both molecular conformation and internal dynamics are affected upon covalent binding of an oligosaccharide to a protein or lipid. Thus, appropriate caution must be exercised when extrapolating from NMR-derived conformation and dynamics results on isolated oligosaccharides to judgments about similar structures linked to proteins or lipids. NMR efforts are getting under way to examine the conformation and dynamics of oligosaccharides as covalent parts of larger molecular weight glycoconjugates (both intact glycoproteins and glycolipids dispersed in lipid bilayers). Amphipatic in nature, isolated glycolipids aggregate in aqueous solution to form micelles. These systems, in defiance of their long overall rotational correlation times, are amenable to high-resolution NMR study and are the most revealing probes available so far for the study of internal motions in lipid-linked oligosaccharides. The conformation and dynamics have been studied of the carbohydrate headgroups of a number of glycolipids inserted in deuterated dodecylphosphocholine (DPC) micelles in DaO and/or H20 [37-40]. As an example, we present here some of our work on ganglioside GDla Neu5Ac'ot(2-+3)Gal'p~(1-+3)GalpNAc~(1-+4)[Neu5Acot(2-+3)]Galp~(1-+4)Glcp~(1-+ 1)Cer embedded in a perdeuterated DPC micelle in aqueous solution. Without the addition of an organic solvent, 1H NMR studies of GDla in a micellar membrane in H20 at --I~ revealed relatively narrow OH signals, allowing their incorporation in extensive NOE studies. It appeared [40] that the observed interresidue 1H/1H NOE contacts (involving both CH and OH protons) could not be explained by only a single conformation of the GDla hexasaccharide such as the quasi-rigid structures proposed in keeping with 13C T1 data for the GDla free oligosaccharide [41 ] in D20 and glycolipid [42] in DMSO. At least two families of carbohydrate conformers [differing in the relative orientation of the Neu5Ac'ot(2-+3)Gal'13(1-+3)external disaccharide arm; see Fig. 7(a)] must exist in this model membrane system to account for the NOE constraints. The internal dynamics of the oligosaccharide were probed by 1H-detection of the 13C relaxation parameters (T1, Tip, and heteronuclear NOEs) of the GDla hexasaccharide in the micellar system at 13C natural abundance. The results of these measurements are listed in Table 3. The 13C relaxation rates R1 and R2 were converted into the spectral densities J(m) at the pertinent frequencies [40,45-47]. Aided by an independent measurement of the global rotation correlation time 'co of the micellar system (from 31p NMR relaxation measurements on the DPC 31p signal), the spectral density data were interpreted at the molecular level with the Lipari-Szabo model-free approach [43,44]. As illustrated schematically in Figure 7(b), the model-free approach distinguishes the types of molecular motion of an oligosaccharide by their different time scales. The total motion of the molecule is separated into the tumbling of the entire aggregate (with rotation correlation time "Co) modulated by more rapid internal (segmental) motion with characteristic correlation time ('ci) and amplitude expressed by an order parameter ($2). The latter is a measure of internal reorientation (S 2 = 1 in the absence of reorientation; S 2 =0 in the presence of completely isotropic reorientation). The separation of

25

Figure 7. (a) Two conformers of GDla on the DPC micellar membrane, differing in the orientation of the outer disaccharide moiety [40]. (b) Diagram of GDla in a DPC micelle illustrating the independent global and local motion model, the characteristics of which can be described by the Lipari-Szabo equations [43,44]. the motions into distinct regimes is crucial to the success of this method. Thus, the internal dynamics (segmental motions) of carbohydrate molecules in solution can be assessed by NMR spectroscopy if the rearrangements occur much faster (or much slower) than overall molecular tumbling. The internal reorientations in GDla on the micellar surface were found to occur with a rotation correlation time of 0.35 ns. The slow tumbling of the micelle (Mr "" 8 kDa) provided the distinct time scale ('to 2.8 ns), facilitating the discovery of the faster internal motion of a couple of flexible glycosidic linkages in the carbohydrate headgroup [40]. The order parameter S 2 was measured for reorientation of the C-H vectors of each glycosyl residue. As computed from 13C Tip, S 2 was found to be --1.0 for internal residues and--0.5 for the terminal (Neu5Ac') and penultimate (Gal') residues most distal to the lipid (Table 3).

26 Table 3. Average 13C relaxation rates a and dynamics parameters b for the headgroup hexasaccharide in Gnla/DPC micelles in aqueous solution Residue

R 1 [s-1]

GalNAc Neu5Ac Gal, Glc

}

'cNeu5Ac' Gal

}

R 2 [s -1 ]

NOE

% [ns]

'17i [ns]

S2 1

2.2+0.2

12.5+2.0

1.2-+0.2

2.8-+0.1

0

2.3+0.2

7.1_+1.0

1.6_+0.2

2.8_+0.1

0.34_+0.10 0.55+0.08

a The 13C relaxation rates are defined as: R 1 = (T1)-1, and R 2 = (T19)-l. b Obtained by nonlinear least-squares optimization using the equations in Fig. 7(b). The relaxation data for the methylene carbons were not included in the analysis. c Data for Gal' C4 were not included.

Application of the methods discussed to larger complex oligosaccharides as well as to those tethered to proteins (see, for example, [48]) or lipid membranes will continue to provide valuable evidence on the question of internal motions on a time scale of the reciprocal of the NMR frequency range used in the experiment (0.1 to 1 ns). Motions on a slower time scale, which would not be detectable by ordinary 13C T1 experiments, can be studied by rotating-frame relaxation data (T10). With control of the spin-locking field strength, the latter parameter can reveal motions on the Bs time scale.

5. CONCLUSIONS Carbohydrate NMR spectroscopy has reached a level of sophistication at which the number of detectable constraints (including hydrogen bonds and glycosidic bond torsion angles) is sometimes large enough to be incompatible with a single rigid conformer, prompting the consideration of ensemble average models. Internal motion in oligosaccharides can be detected and quantified by NMR spectroscopy if it occurs relatively fast compared to the overall motion of the molecule. Tethering the oligosaccharide to a phospholipid micelle in aqueous solution satisfies this criterion. Knowledge of three-dimensional conformations and the dynamics/flexibility of complex carbohydrates in their natural environment will broaden our insight into their functioning as mediators of numerous biological cell-cell and cell-molecule interactions. It is anticipated that NMR spectroscopy will continue to make invaluable contributions towards this goal.

6. ACKNOWLEDGMENTS The authors thank Drs. Leszek Poppe and John Glushka for helpful discussions and Rosemary Nuri for editing the manuscript. This research is supported by National Institutes of Health (NIH) grant P41-RR-05351 and Department of Energy (DOE) Plant Science Center grant DE-FG09-93ER20097.

27 7. A B B R E V I A T I O N S

The abbreviations used are: Cer, ceramide; COSY, homonuclear scalar correlation spectroscopy; DPC, dodecylphosphocholine; f, furanose; HMBC, heteronuclear multiplebond connectivity; HMQC, heteronuclear multiple quantum coherence; HSMBC, heteronuclear single- and multiple-bond connectivity; HSQC, heteronuclear single quantum coherence; MD, molecular dynamics; n-D, n-dimensional (where n=l, 2, or 3); NMR, nuclear magnetic resonance; NOE, nuclear Overhauser effect; NOESY, NOE correlated spectroscopy; p, pyranose; ROESY, rotating-frame Overhauser and exchange spectroscopy; SL6, sialyl-o~(2~6)-lactose; TOCSY, total scalar correlation spectroscopy. The symbols used are defined as follows: J, scalar coupling constant; J(c0), spectral density at frequency co; r, distance; S2, generalized order parameter; c~, cross-relaxation rate; Tt, longitudinal relaxation time; T2, transverse relaxation time; T10, longitudinal relaxation time in the rotating frame; "Co, global rotation correlation time; xi, internal (local) rotation correlation time; ~), torsion angle H1-C1-O1-C'x; ~t, torsion angle C1-O1-C'x-H'x; o3, torsion angle H5-C5-C6-O6.

8. REFERENCES

1 H. van Halbeek, in "Encyclopedia of Nuclear Magnetic Resonance," D.M. Grant and R.K. Harris (eds.), Wiley, Chichester, 1995, in press. 2 J. Dabrowski, in "Two-Dimensional NMR Spectroscopy. Applications for Chemists and Biochemists (Second Edition)," W.R. Croasmun and R.M.K. Carlson (eds.), VCH, New York, 1994, p. 741. 3 S.W. Homans, Prog. NMR Spectrosc., 22 (1990) 55. 4 S.W. Homans, in "Molecular Glycobiology," M. Fukuda and O. Hindsgaul (eds.), Oxford University Press, Oxford, 1994, p. 230. 5 S. PErez, A. Imberty, and J.P. Carver, Adv. Comput. Biol., 1 (1994) 147. 6 H. van Halbeek, Curr. Opin. Struct. Biol., 4 (1994) 697. 7 M.C.R. Symons, J.A. Benbow, and J.M. Harvey, Carbohydr. Res., 83 (1980) 9. 8 J. Dabrowski and L. Poppe, J. Am. Chem. Soc., 111 (1989) 1510. 9 L. Poppe and H. van Halbeek, J. Am. Chem. Soc., 113 (1991) 363. 10 L. Poppe and H. van Halbeek, Nature Struct. Biol., 1 (1994) 215. 11 B. Adams and L.E. Lerner, Magn. Reson. Chem., 32 (1994) 225. 12 J.C. Christofides and D.B. Davies, J. Am. Chem. Soc., 105 (1983) 5099. 13 V. Sklenfir and A. Bax, J. Magn. Reson., 74 (1987) 469. 14 V. Sklenfir, R. Tschudin, and A. Bax, J. Magn. Reson., 75 (1987) 352. 15 V. Sklenfir and A. Bax, J. Magn. Reson., 75 (1987) 378. 16 M. GuEron, P. Plateau, and M. Decorps, Prog. NMR Spectrosc., 23 (1991) 135. 17 B. Mulloy, T.A. Frenkiel, and D.B. Davies, Carbohydr. Res., 184 (1988) 39. 18 V.H. Tran and J.W. Brady, Biopolymers, 29 (1990) 961. 19 V.H. Tran and J.W. Brady, Biopolymers, 29 (1990) 977. 20 C. Herv6 du Penhoat, A. Imberty, N. Roques, V. Michon, J. Mentech, G. Descotes, and S. PErez, J. Am. Chem. Soc., 113 (1991) 3720. 21 J.M. Duker and A.S. Serianni, Carbohydr. Res., 249 (1993) 281. 22 G.M. Brown and H.A. Levy, Acta Crystallogr., Sect. B, 29 (1973) 790. 23 D.C. McCain and J.L. Markley, Carbohydr. Res., 152 (1986) 73. 24 D.C. McCain and J.L. Markley, J. Am. Chem. Soc., 108 (1986) 4259.

28 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48

D.C. McCain and J.L. Markley, J. Magn. Reson., 73 (1987) 244. L. Poppe and H. van Halbeek, J. Am. Chem. Soc., 114 (1992) 1092. D.B. Davies and J.C. Christofides, Carbohydr. Res., 163 (1987) 269. B. Adams and L. Lerner, J. Am. Chem. Soc., 114 (1992) 4827. A. Bax and M.F. Summers, J. Am. Chem. Soc., 108 (1986) 2093. F.J. Cassels and H. van Halbeek, Methods Enzymol., 253 (1995) 69. A. Bax and D. Marion, J. Magn. Reson., 78 (1988) 186. J.J. Titman, D. Neuhaus, and J. Keeler, J. Magn. Reson., 85 (1989) 111. G. Zhu and A. Bax, J. Magn. Reson. Ser. A, 104 (1993) 353. G. Zhu, A. Renwick, and A. Bax, J. Magn. Reson. Ser. A, 110 (1994) 257. P. Huber, C. Zwahlen, S.J.F. Vincent, and G. Bodenhausen, J. Magn. Reson. Ser. A, 103 (1993) 118. L. Poppe, R. Stuike-Prill, B. Meyer, and H. van Halbeek, J. Biomol. NMR, 2 (1992) 109. L. Poppe, C.-W. vonder Lieth, and J. Dabrowski, J. Am. Chem. Soc., 112 (1990) 7762. D. Acquotti, L. Poppe, J. Dabrowski, C.-W. von der Lieth, S. Sonnino, and G. Tettamanti, J. Am. Chem. Soc., 112 (1990) 7772. H.-C. Siebert, G. Reuter, R. Schauer, C.-W. vonder Lieth, and J. Dabrowski, Biochemistry, 31 (1992) 6962. L. Poppe, H. van Halbeek, D. Acquotti, and S. Sonnino, Biophys. J., 66 (1994) 1642. S. Sabesan, J.O. Duus, T. Fukunaga, K. Bock, and S. Ludvigsen, J. Am. Chem. Soc., 113 (1991) 3236. J.N. Scarsdale, J.H. Prestegard, and R.K. Yu, Biochemistry, 29 (1990) 9843. G. Lipari and A. Szabo, J. Am. Chem. Soc., 104 (1982) 4546. G. Lipari and A. Szabo, J. Am. Chem. Soc., 104 (1982) 4559. S. Bagley, H. Kovacs, J. Kowalewski, and G. Widmalm, Magn. Reson. Chem., 30 (1992) 733. P.J. Hajduk, D.A. Horita, and L.E. Lerner, J. Am. Chem. Soc., 115 (1993) 9196. J. Kowalewski and G. Widmalm, J. Phys. Chem., 98 (1994) 28. T.J. Rutherford, J. Partridge, C.T. Weller, and S.W. Homans, Biochemistry, 32 (1993) 12715.

S.B. Petersen, B. Svensson, and S. Pedersen (Eds), CarbohydrateBioengineering 9 Elsevier Science B.V. All rights reserved.

29

Linkage analysis by mass spectrometry of chemically modified oligosaccharides from glycosphingolipids and glycoproteins Bo Nilsson National Defence Research Establishment, Department of NBC Defence, S-901 82 Ume~t, Sweden

Abstract Mass spectrometry of oligosaccharides gives structural information on monosaccharide sequence and depending on ionisation technique used, molecular weight. In order to extend the structural information to also include glycosidic linkage positions, specific chemical modif'lcations were introduced. Trifluoroacetolysis was used to specifically degrade high-mannose structures from the reducing terminal. Periodate oxidation followed by reduction, peracetylation or permethylation and analysis of the products by mass spectrometry gave on the basis of the sequence ions linkage positions between monosaccharide residues in oligosaccharides obtained from glycosphingolipids and glycoproteins.

1. INTRODUCTION In order to understand how biological compounds interact i.e. the structure - function relationship, structural analysis is essential. Biological phenomena work through specific interactions of chemical structures. Recent developments of cell systems for manufacturing of biologically important proteins for use in therapy have focused on post translational modifications such as glycosylation. Structural analysis of the carbohydrate chains has therefore become important in order to avoid cell systems that produce glycoproteins with glycosylation, which in humans can cause undesired immunological reactions and unfavourable serum clearance rate. Another type of biologically important glycoconjugates are glycosphingolipids, which are cell surface associated. Since the carbohydrate portion is exposed to the exterior it should be accessible to interactions with other molecules. Neutral and acidic glycosphingolipids have been implicated in several roles, for example receptors for bacteria, virus, bacterial toxins and as tumour associated antigens. Analytical methods for proteins are well developed, but carbohydrate-containing polymers have for long time been difficult to analyse. One of the reasons is that no other biological compounds can form so many isomeric species as carbohydrates. The great number of compounds that can be formed also indicates that the content of biological information is enormous. The biological functions of glycoconjugates are however still not completely

30 understood. Carbohydrates have in the past, in contrast to proteins and nucleic acids, been regarded as less interesting from biological point of view, which have resulted in that less efforts have been investigated in analytical techniques for this type of compounds. When spectroscopic methods as mass spectrometry (MS) and NMR were introduced a dramatic progress in structural analysis of glycoconjugates was noticed. A comprehensive strategy including MS and NMR for glycoprotein analysis has previously been published [1]. This contribution will focus on recent developments in mass spectrometry of glycosphingolipid- and glycoprotein oligosaccharides.

2. MASS SPECTROMETRY Structural analysis of glycoconjugates usually involves various types of mass spectrometric techniques. Traditionally electron ionisation mass spectrometry (EI-MS) of derivatised monosaccharides and oligosaccharides has been used. Compounds have to be volatile in order to be analysed by this ionisation technique. When fast atom bombardment mass spectrometry (FAB-MS) was introduced underivatised as well as derivatised high molecular weight glycoconjugates could be analysed. Later introduced techniques such as electrospray mass spectrometry (ES-MS) and time of flight mass spectrometry (TOF-MS) have further extended the mass range, allowing molecular weight determination of intact glycoproteins. These recently developed techniques have focused mainly on molecular weight determination rather than structural analysis. Carbohydrate-containing polymers differ from other biological substances in features of branching, multiple binding positions and the monosaccharide residues can usually not be discriminated by their masses. Structural analysis of glycoconjugates by mass spectrometry is therefore a special challenge. The structural information, that can be obtained, includes molecular weight and monosaccharide sequence in terms of hexose, deoxyhexose acetamido-deoxyhexose and sialic acid. In order to completely determine a structure several other features have to be assigned e.g. linkage positions between monosaccharide residues. Mass spectra are often recorded after derivatisation, peracetylation or permethylation, which in FAB-MS enhances the sensitivity. The sequence ions of derivatised compounds are formed by ionisation of the ring oxygen followed by a homolytic cleavage process of glycosidic bonds yielding cyclic oxonium ions. Presence of branch points can be determined by the absence of certain sequence ions. Only in exceptional cases binding positions between monosaccharides can be determined e.g. substitution positions of permethylated 2-acetamido-2-deoxyhexosyl (HexNAc) residues. After a primary cleavage of the HexNAc glycosidic bond in EI-MS or FAB-MS the relative intensities of the secondary fragments formed by eliminations determine the substitution pattern (Figure 1). The elimination takes place preferentially from the 3-position of the HexNAc residue [2]. Substitution positions of monosaccharide residues can be determined by gas chromatography-mass spectrometry (GC-MS) of partially methylated alditolacetates [3], but the sequential order of linkages is in general difficult to determine. NMR-Analysis or enzymatic methods can often provide this information if sufficient material is available. In order to extend the structural information to include linkage analysis in oligosaccharides by mass spectrometry, specific chemical modifications were introduced.

31 228

%

432

%

Hexl --4HexNAc I -

Hexl-3HexNAcl-

464 464

!

196

196 228

I .

m/z 402

m/z 432

% Hexl-3HexNAcl4 I deoxyHexl

196

I,

Hex ! -4HexNAe

638

4?2

I

I-

3 I deoxyHexl 638

196 ,

,

402 [

m~

Figure 1. Relative intensities in FAB-MS of primary and secondary ions derived from 3- and 4mono- and 3,4-disubstituted permethylated HexNAc residues.

3. CHEMICAL MODIFICATIONS Periodate oxidation was used to specifically modify monosaccharide residues in oligosaccharides [4]. Bonds connecting carbons carrying hydroxyl groups in vicinal positions are oxidatively cleaved and aldehydes are formed, which subsequently are reduced with NaBD4 (Figure 2). The product is then analysed after peracetylation or permethylation by GC-MS or FAB-MS depending on the size of the oligosaccharide. I periodate HCOH oxidation

I / C,,, NaBD 4 I acetylation or ] H O reduction HDCOH methylation HDCOAc

HCOH

H,

I

"c

I

//O

HDCOH

t

HDCOAc

I

I

HDCOMe or

HDCOMe I

Figure 2. Reaction steps used for preparation of derivatives for analysis by mass spectrometry.

The products obtained depend on linkage positions between residues and the mass values of ions formed by cleavage of glycosidic bonds for terminal and internal residues after permethylation and peracetylation are shown in Table 1.

32 Table 1 Mass values for terminal and intemal residues after periodate oxidation, NaBD4-reduction and derivatisation Residue

Permethylation

Peracetylation

Non-reducing Hexl6-deoxyHexlHexNAclNeu5Ac2-

179 149 264 289

263 205 334 a

208 204 208 164 245 245

292 288 292 220 287 287

Internal -2Hexl-3Hexl-4Hexl-6Hex 1-3HexNAcl-4HexNAc 1-

Residue

Permethylation Peracetylation

-6HexNAcl-8Neu5Ac2-

249 361

291 a

105 192 148 104

161 304 232 160

-3HexNAc-ol 189 -4HexNAc-ol 233 -6HexNAc-ol 60

231 303 88

Reducing -2Hex -3Hex -4Hex -6Hex

Reduced

a No mass due to lactone formation

Terminal residues are always oxidised and internal hexosyl (Hex) residues are oxidised if 2-, 4- or 6-substituted, whereas 3-substituted are not. For permethylated compounds 2- and 4substituted Hex residues both give a mass increment of 208 mass units (m.u.), but the structures are different. A secondary fragment, formed by elimination of methanol, is seen from the 2-substituted, but not for the 4-substituted Hex residue [5]. In peracetylated samples these two products give in addition an ion formed by cleavage of the bond between carbon 5 and the former ring oxygen, which for the 2-substituted residue is constant of m/z 160, whereas for the 4-substituted the m/z value depends on the mass of the substituent [6]. Both 3- and 4substituted HexNAc residues are resistant to periodate oxidation. As in unoxidised and permethylated compounds these residues are differentiated by the secondary fragments formed by eliminations as will be illustrated by examples below. In order to determine the substitution position of a reducing 2-acetamido-2-deoxyhexose the periodate oxidation has to be carried out on the reduced compound. Substitution positions of branched residues can sometimes be determined (Table 2). A 3,6-disubstituted Hex-ol gives after periodate oxidation and NaBD4-reduction two products, ethylene glycol and glycerol. O-Linked glycans often contain a 3,6-disubstituted HexNAc-ol (GalNAc-ol), which will be cleaved by periodate giving two products, ethylene glycol and N-acetyl tetrosaminitol. In N-linked structures the HexNAc-ol (GlcNAc-ol) is usually 4,6-disubstituted, which is not cleaved by periodate. Another chemical modification, trifluoroacetolysis, is carried out in a mixture of trifluoroacetic acid (TFA) and trifluoroacetic anhydride (TFAA). This mixture has a powerful

33 acetylating property resulting in rapid trifluoroacetylation of hydroxyl groups. Glycosidic bonds are stabilised by inductive effect exerted by the O-trifluoroacetyl groups [7].

Table 2 Products obtained from some common reduced branched residues after periodate oxidation and NaBD4-reduction Branched residue Products

CHDOH 6

Hex-ol

/3

CHDOH

!%

,~

I CH [

CHDOH

~I~OH

CHNHAc ~6

3 HexNAc-ol

CHDOH [

c%

I I CHDOH

~CH

~HaOH

CHNHAc

I I ~CH I CHOH I ~Ct-I 2

CHOH

~6

/

4

HexNAc-ol

Oligosaccharides, N- and O-linked, are released from glycoproteins, as N-trifluoroacetylated derivatives [8, 9]. Under certain conditions glycoprotein oligosaccharides are specifically degraded from the reducing terminal [ 10]. Trifluoroacetolysis is also useful for liberation of the carbohydrate portion from glycosphingolipids [11]. Periodate oxidation in combination with trifluoroacetolysis has been used in structural analysis of high-mannose glycoprotein oligosaccharides, as will be discussed below.

34 4. GLYCOSPHINGOLIPIDS Neutral and acidic glycosphingolipids, isolated various species and cell types, have been subjected to numerous studies of their biological functions. Due to their lipophilic nature these types of compounds are suitable for separation by thin layer chromatography, which has been used for identification by comparison of migration with reference compounds. Neutral glycosphingolipids and gangliosides can be analysed by FAB-MS as underivatised compounds or after permethylation. Besides monosaccharide sequence, composition of the ceramide moiety can be deduced from the spectra [12]. Only in permethylated samples and in the presence of internal HexNAc residues substitution position of these residues can be determined as previously discussed. In order to determine other glycosidic linkages by FAB-MS, periodate oxidation can be used on intact glycosphingolipids [5]. An alternative approach is to release the oligosaccharide from the ceramide portion and then perform the periodate oxidation.

4.1. Release of oligosaccharides The carbohydrate portion of glycosphingolipids can be released by chemical or enzymatic methods. Treatment of neutral glycolipids or gangliosides, containing an unsaturated sphingosine base, with ozone followed by mild base liberates the oligosaccharide [ 13]. Another chemical method, trifluoroacetolysis, also requires an unsaturated sphingosine base and works best on neutral glycosphingolipids. When used on gangliosides partial loss of sialic acid is seen. From N-acetyl hexosamine-containing glycosphingolipids the oligosaccharides are released as N-trifluoroacetyl derivatives, which after permethylation are suitable for analysis by GC-MS [ 14]. An altemative to chemical methods is treatment with enzymes. Enzymatic methods have an advantage in that both the carbohydrate moiety and the ceramide can be recovered separately, which is useful in studies of the ceramide residue. There are number of ceramidases commercially available e. g. from the medicinal leech and from the earth worm Lumbricus terrestris [ 15, 16]. 4.2. Mass spectrometry of glycosphingolipid oligosaccharides Ozonolysis of the GM3 ganglioside released the oligosaccharide: Neu5Acot2-3Gall] 1-4Glc The trisaccharide was subjected to periodate oxidation, NaBD4-reduction and permethylation. Analysis of the product by GC-MS gave a spectrum shown in Figure 3. The primary sequence ion of m/z 289 together with a secondary of m/z 257, formed from the primary by elimination of methanol, are characteristic for a periodate oxidised non-reducing terminal sialic acid. A disaccharide sequence of Neu5Ac2-3Gall- is deduced from the above ions together with rn/z 493. The linkage position between these residues is determined by the mass increment of 204 m.u. representing a periodate resistant hexosyl residue, which therefore must be 3-substituted (Table 1). The alditol-containing ions of m/z 148 and 352 show that the reducing glucose is 4-substituted. These two ions give rise to secondary ions of m/z 116 and m/z 320, respectively, by elimination of methanol. An ion of m/z 89, formed by cleavage within the alditol, shows two O-methylated carbons. El-Mass spectra of carbohydrates never give molecular ion species. In this case an [M-59] § ion of m/z 598 is seen, which is a loss of a

35 methoxycarbonyl radial (. COOMe) from the molecular ion, typical for EI-spectra of sialic acid containing oligosaccharides.

257

188. 9B

289

w r 8B z "~ 7B o z 6B =)

Me

~.

181

LU 4B >

3B

~ 2B w n- IB

~N

31

89

493

MeOOC--I]

Mel

co 5B

~

257+- 289 MeO

i

I

I

L--'0"+" 352

I

ICHOMe

I CIH20Me 148 [M-59]+

493

1481 183

......... t,++

CHDOMe

O

#' "-I-- I ~

159 I

116

ICH2OMe--7

,. . 2++

. . . . . . . . . . . . . . . . . .

,,,

4++

3++

....

1.4!946,5..I,

[ +B+

598 . . . . . . . . . . .

m/z

+m+

Figure 3. GC-MS spectrum of the GM3 oligosaccharide after periodate oxidation, NaBD4reduction and permethylation.

Treatment of the GTlb ganglioside with ozone liberated the oligosaccharide with the structure of:

Neu5Ac0~2-3Gal[31-3GalNAc[~ 1 ~ 4 Neu5Ac~176

j

3 Gal131-4Glc

A FAB-mass spectrum obtained after the above periodate treatment is shown in Figure 4. A disialyl sequence of Neu5Ac2-8Neu5Ac2- can be deduced from the primary sequence ions of m/z 289 and m/z 650, where the latter ion is an increase of 361 m.u. (Table 1). The linkage between these residues must therefore be 2-8 since the intemal sialic acid residue is not oxidised by periodate. Any other linkage position between these residues would make the internal sialic acid susceptible to periodate and thereby give rise to other ions than the above. Another sialylated sequence of Neu5Ac2-3Gal 1- is determined by the primary sequence ions of m/z 289 and m/z 493 as previously discussed. This monosialylated sequence is linked to the 3position of the GalNAc, determined from the primary sequence ion of m/z 738 and a secondary of m/z 228. Elimination of the Neu5Ac2-3Gal sequence gives rise to m/z 228 specific for a 3substituted 2-acetamido-2-deoxyhexosyl residue [ 17]. The substitution pattern of the branched

36 galactosyl residue cannot be inferred from the spectrum. An [M+23] + ion (molecular ion plus sodium) of m/z 1765 is consistent with a periodate oxidised G-m, oligosaccharide. After trifluoroacetolysis of globoside and reconstitution to the N-acetyl derivative the globo-N-tetraose oligosaccharide was obtained: GalNAc[31-3Galc~ 1-4Gal]31-4Glc

289 228

21i7

[M+23]* 17 ;5

,u ;8. z0

~ 58. m ~ 58.

g,

]8.

1888

m/z

493

288

488

]88

738

588

257 ..,-289 MeOOC'~

G88

493 CH2OMe~

228 "4---738 CH2OMe" ~

1

MeO

I

MeO MeOOC I

Me ~ .

,o%,---od Meo

~

257 4-- 289

788

MeNAc Me ~.

MeOOC

I

,,c-'~,~--~od MeO

___1

888

CH2OMe

/

MeO

2

988

1~8

m/z

?HDOMe

CH2OMe

650

Figure 4. FAB-MS spectrum of the G'rlb oligosaccharide after periodate oxidation, NaBD4reduction and permethylation.

Treatment with periodate as above and analysis by FAB-MS gave a spectrum shown in Figure 5. A periodate oxidised non-reducing terminal GalNAc is determined by the primary

37 sequence ion of m/z 264 and a secondary of m/z 232, formed by elimination of methanol. These ions combined with m/z 468 and m/z 676 show a sequence of GalNAcl-3Gall-4Gall-. The ion of rn/z 676 is an increase of 208 m.u. to m/z 468 and means a 2-substituted or 4-substituted Hex residue. Substitution in the 2-position can however be excluded since no secondary fragment, formed by elimination of methanol, is seen [5]. Substitution of the reducing glucose in the 4-position is calculated from the [M+23] + ion of m/z 863 and the sequence ion of m/z 676.

264

CH,,OMe

IJJ (J z I-

MeODHC

I

676

CH,~OMe'~

~,HoOMe'~

]~l' J, " ~ m ~ M e O D H C OMe

CHDOMe

CHDOMe

MeNAc

i HO CH2OMe

68

""< ..J UJ

2(i4

flail. 96. 88 ~0

468

~t t,

>400 x3 it

232

4g.

31.

1s2 let 14e

I

I

347

468

'[

I

7~9 I

~

I

,[

J~s

e37863 IM+2 m/z

m

Figure 5. FAB-MS spectrum of globo-N-tetraose after periodate oxidation, NaBD4-reduction and permethylation. Reprinted from: Carbohydr. Res. 168 (1987) 15. Copyright 1995 Elsevier.

5. GLYCOPROTEINS Production of recombinant proteins and monoclonal antibodies for use in therapy and diagnosis has renewed the importance of glycoprotein analysis. A complicating factor when working on glycoproteins is the heterogeneity among the oligosaccharides. A given glycosylation site often has several different structures. In order to carry out structural analysis the oligosaccharides have to be released from the polypeptide chain and purified to homogeneity. Separation of released oligosaccharides has traditionally been a problem due to the small differences in structure. New HPLC columns have, however, dramatically improved the separation methods, allowing separation of neutral as well as acidic isomeric structures [ 18, 19]. Structural analysis has been carried out by chemical, enzymatic and spectroscopic methods. Complete structural determination can in most cases be performed by NMR

38 spectroscopy if sufficient material is available. Mass spectrometry usually requires less material but complete structural determination cannot be obtained. 5.1. Release of oligosaccharides There are a number of chemical and enzymatic methods to liberate carbohydrates from glycoproteins. A chemical method frequently used to release N-linked oligosaccharides is hydrazinolysis [20]. After N-acetylation and mild acid treatment the oligosaccharides are recovered as free reducing compounds. O-Linked oligosaccharides are partially degraded from the reducing terminal, using this method. The most commonly used method to release O-linked glycans is reductive 13-elimination [21 ]. By using harsher conditions this procedure can also be used to liberate N-linked structures, which then have to be re-N-acetylated [22, 23]. When using these alkaline degradation methods the carbohydrates are recovered as oligosaccharide alditols. Trifluoroacetolysis is another chemical method for release of N- and O-linked glycans [8, 9]. Sialic acid is partially lost, which makes this procedure more suitable for asialoglycoproteins. The fate of a reducing hexosamine depends on the proportions of TFA/TFAA and under certain conditions this method can be used to specifically modify high-mannose structures, as will be discussed below [24]. All chemical methods suffer more or less from degradations or modifications of the reducing terminal of the released glycans. A viable alternative is therefore enzymatic methods. Several specific endoglycosidases are commercially available. Intact N-linked oligosaccharides can be obtained by treatment with PNGase F from Flavobacterium meningosepticum. For release of O-linked glycans there is an O-glycanase which releases the disaccharide GalI31-3GalNAc from Ser/Thr but not if this disaccharide is substituted. The specificity and sources of endoglycosidases have been reviewed elsewhere [25]. After separation of released glycans structural analysis can be carried out. 5.2. Mass spectrometry of N-linked oligosaccharides Glycoprotein oligosaccharides have been subjected to structural analysis by FAB-MS mostly as permethylated derivatives. In order to extend the structural information beyond sequence and molecular weight, periodate oxidation was used. The procedure used includes the same reaction steps as previously mentioned for glycosphingolipid oligosaccharides. Since almost all glycoprotein oligosaccharides, N- or O-linked, contain a reducing terminal HexNAc the periodate oxidation has to be carried out on the NaBIL-reduced compound in order to be able to determine the substitution position of this residue (Tables 1 and 2). The above procedure was applied to a biantennary oligosaccharide alditol with a structure of: Neu5 Aco~2-6G al [31-4GlcNAc 131-2Manor1 6 /3 Neu5 Ac~2- 6G all31-4GlcNAc]31-2Manor 1"

Man[31-4GlcNAc]31-4GlcNAc-ol

The FAB-mass spectrum obtained is shown in Figure 6. A non-reducing terminal periodate oxidised sialic acid is recognised by the primary and secondary fragments of m/z 289 and m/z 257. These sequence ions combined with m/z 698, from which a secondary ion of m/z 666 is formed, determine a sequence of Neu5Ac2-6Gall-

39 4GlcNAcl-. As previously discussed elimination from a HexNAc takes preferentially place from the 3-position in this case elimination of methanol giving m/z 666 [ 17]. Substitution in the 6-position of the GlcNAc residue can be excluded since this residue would then be oxidised, 289 U..I

Z

2520

9g

80 7g

Z

257

G6

<1: LU Sg i-'-

4it

_J LU 3g,

228 L 233

High mass scan

2gg

378

3oo

478

48g

378 (453)

6e~. . . . . . . . .

seB

o.IIo/, o, ~'CHDOMe ~J I / J' ~ P "" "~_j~'MeOCHD / Iu\ ~ i MeODHC' I "

MeO

CH2OMe~ ] o

J ~

378 MeOOC O

/

I

257 ~-- 289

MOeocHD

l

874 4--906

o

e

233

~ Hoh

o

~MeOL~

MeODHC r

I

(453)

O.

OMe

~ ~

CHrOMe

O

O

e

O

~]

MeO ~

, ..e N ~Ac

478

J--'--CH2OMe j CHOMe

MeODHC ==m=~If~,JrO --CH2

CHrOMe O

7a. . . . . . . . . 0h. . . . . . . . . sa. . . . . . . . i0'D8

CH2OMe"~

MeNAc

I - CH'~

874 906 m/z

666 ~ - 698

MeOOC---] f- CH-~2 ~

CHDOM.

666

Low mass scan

257 4-- 289

>800 x3

I

ii i

~L~

Me.

m/z 698

196

2g.

Ac/

[M+23]*

OO M ~ ~ O T '

~H I CHDOMe

MeNAc

I M e O D H C ==~====If "I"*O

666 ~-- 698

874 ~-- 906

Figure 6. FAB-MS spectrum of a biantennary oligosaccharide alditol after periodate oxidation, NaBD4-reduction and permethylation. Reprinted from: Protein Glycosylation, Cellular, Biotechnological and Analytical Aspects. GBF Monographs 15 (1990) 125. Copyright 1995 VCH.

giving other sequence ions than the above. No sequence ion corresponding to cleavage of the oxidised Gal residue, which would be m/z 453 is seen, instead an ion of m/z 378 is formed by cleavage of the carbon 5 and the former ring oxygen bond of the periodate degraded Gal residue. Both antennae of this biantennary structure contain 2-6 sialylated sequences. These sequences are linked to the 2-position of a mannose residue, deduced from the primary and secondary ions of m/z 906 and m/z 874, respectively. As previously discussed a periodate oxidised 2-substituted hexosyl residue gives a secondary ion by elimination of methanol, in this case m/z 874, which is not seen for a 4-substituted (Figure 5). Using the periodate oxidation

40 allows determination of all glycosidic linkage positions in the linear sequences. Substitution positions of the branched mannose residue can, however, not be determined from the mass spectrum. Since the periodate oxidation was carried out on the reduced compound the 4substitution of the reduced GlcNAc residue is determined from the ion of m/z 233 and combined with m/z 478 the chitobiose structure is established. An [M+23] § ion of m/z 2520 gives supporting evidence for the structure. This method is also useful when screening for sequences in mixtures of structures. The triantennary oligosaccharide fraction from a mouse monoclonal IgA antibody was analysed by FAB-MS after the above periodate treatment [26]. The sequence ions determining linkages in the outer linear sequences are indicated in Figure 7.

289

Gal IIO--4GIcNAcl.-]-O I 392--424 GalIIOB3GaI1--O--4GIcNAcl-~.

88. u.I

r~ Z

<

J

596"- 628

78.

cl z::::) 61t. m ,< 58, LIJ > i
257

257-'-

289

378, (453)

666-- 698

Neu5Ac2-]--OI3G.~O~4GIcNAc_~-O~ 257"-289

493

706"-738 Fucl

196

I

6

BO~lcNAc--ol

I 214

410 ~TR 3 9 2 424 -;~ \ 41 01

288

388

488

628 I

fin

688

698 I I 706

7ffi

m/z

8811

Figure 7. Part of a FAB-MS spectrum of a mixture of triantennary structures after periodate oxidation, NaBD4-reduction and permethylation. Reprinted from: Arch. Biochem. Biophys. 300 (1993) 335. Copyright 1995 Academic Press.

All GlcNAc residues in the antennea of the triantennary structures are 4-substituted deduced from the secondary fragments of m/z 392, m/z 596, m/z 666 and m/z 706, which all are results of eliminations of methanol. Furthermore the ion of m/z 410 shows that the reducing GlcNAc is substituted in the 6-position by a fucose residue. High-mannose structures are less favourable to analyse by FAB-MS as permethylated derivatives due to the absence of branches with N-acetyl hexosamine residues, which as shown above give abundant sequence ions. Peracetylated derivatives are in this case preferable. Two

41 methods for linkage analysis of high-mannose structures by mass spectrometry have previously been published [6, 24]. The first method is based on periodate oxidation and analysis by FABMS and EI-MS of peracetylated and permethylated derivatives. This method allows determination of all glycosidic linkage positions including substitution of branched residues. An example of application of this method is demonstrated on a Man6GlcNAc compound with the following structure: Manor 1 ~ 6 Manor 17

3 Manc~1 ~ 6

Manor 1-2Mano~1j

Man~31-4GlcNAc

3

Periodate oxidation of the NaBH4-reduced compound followed by NaBD4-reduction, permethylation and analysis by FAB-MS gave a spectrum shown in Figure 8. An intense sequence ion of m/z 179 is seen representing the non-reducing terminal mannose residues. A secondary sequence ion of m/z 355 formed by elimination of methanol from m/z 387, which is not seen, determines the Manl-2Manl- sequence [6]. Substitution of the reducing GlcNAc residue in the 4-position is established by the ion of m/z 233. The spectrum also gives information on substitution positions of branched residues. Studies on high-mannose structures have shown that 3,6-disubstituted residues give rise to alditol-containing ions where the positive charge is located on carbon 6 of the disubstituted residue [6]. Two such ions are seen in the spectrum, m/z 809 and m/z 1177. The former ion shows that the Man 1-2Man 1- sequence is linked to the 3-position of the 3,6-disubstituted Man adjacent to the reducing GlcNAc residue. The latter ion, m/z 1177, shows that the other 3,6-disubstituted residue is substituted in the 3-position by a single Man residue. From the [M+I] + ion of m/z 1373 it can be calculated that this branched residue also carries a single Man residue in the 6-position. The intensifies of the branch-specific ions of m/z 809 and m/z 1177 are more prevalent in direct-probe EI-MS. The second method, which is exemplified below, uses degradation with trifluoroacetolysis. Treatment with TFA/TFAA in proportions 1/1 (v/v) at 100~ for 48 h specifically degrades compounds with a reducing terminal N-acetyl hexosamine resulting in products quantitatively depleted of the reducing HexNAc residue [ 10]. For high-mannose structures, as protein-linked or as free reducing compounds, this means that the chitobiose sequence is lost and a mannose residue occupies the reducing terminal position. High-mannose compounds frequently contain a 3,6-disubstituted residue linked to the chitobiose. After TFA/TFAA treatment and reduction (NaBI-h) periodate oxidation is carried out followed by a second reduction (NaBD4). The 3,6disubstituted mannitol will be cleaved by periodate giving two products a glycerol containing two deuteriums and an ethylene glycol with one deuterium (Table 2). The products obtained are analysed after peracetylation by FAB-MS or GC-MS. The procedure described above was applied to the following pentasaccharide: Manotl-3Manotl~ 6 Manor 1j

3 Manl31-4GlcNAc

42 The GC-MS spectra obtained are shown in Figure 9.

IW.

233

179

1177

~._c~ICH2OMe | i

1' '9

MeODHCMeoDHC" "I- O .CH2

809

179 ~MeOOt~ ~'~~~

c.,o,. ] M.O7 / I-2-~

LU 78,

@ z <

M ' ~ 1 7 6 oo. c

68, z no 50.

W ~

='~

" . . . . . . . ___! 179

3O

l

498

1

0

J. CHDOMe

J

3 5 5 - - - (387)

>650 x5

355

o

~

I CH2OMe

[~ .....Ac J~.OM. otc.

CH2OMe t MeO~mm~

Oo. c

MeODHC

o

- o - - -

CH2OMe X

IJJ >~ 4o

~

233

[ M+I]+ 1373 /

_

809

1177

......

1~$2

'LLL ''

m/z

Figure 8. FAB-MS spectrum of Man6GlcNAc-ol after periodate oxidation, NaBD4-reduction and permethylation. Reprinted from: Methods in Molecular Biology: Glycoprotein Analysis in Biomedicine 14 (1993) 35. Copyright 1995 Humana Press. A periodate oxidised, NaBD4-reduced and peracetylated non-reducing terminal hexosyl residue is recognised by the cleavage ions of m/z 160 and m/z 263. The former ion is formed by cleavage of the bond between carbon 5 and the former ring oxygen. Spectrum A shows a glycerol residue from m/z 161 and together with m/z 264, formed by cleavage within the oxidised non-reducing terminal Man residue, it can be concluded that the spectrum represents the mannose linked to the 3-position of the 3,6-disubstituted Man residue in the original pentasaccharide. The other component formed gave spectrum B. An ethylene glycol residue is determined from the ion of m/z 88. The sequence ions of m/z 263 and m/z 551 determine a sequence of Manl-3Manl-. Sequence ions containing the ethylene glycol residue of m/z 376 and combined with m/z 479, formed by cleavage within the oxidised non-reducing terminal, confirm the linkage of the disaccharide sequence. This structure represents the Manl-3Manlbranch, which is linked to the 6-position of the 3,6-disubstituted mannose residue of the

43 original pentasaccharide. Using this method all glycosidic linkage positions including substitution positions of the branched residue can be determined.

uJ () lOe_ < I~1 88 z 23 El 68 < W

160

160 161

A

161 U--

CH '2OAc

I ~

264 / O r / - - - - I CHDOAc

Acoo.:.' ?~_- o1'~ ~coDNC, I " IF 263

CHDOAc

48

> 200 x 5

263 264

--I

"[:I: '

O

..~,L

'~

..........

60

80

d. "~

i1~I.

.... ,,

~08

'I ~

.. I

i~8

. . . . . .

1i8

,'~ II

,~,.~ I. .

168

100

,~,

R16 ,,,,~

L

~8

2~0

2i0

. . . .

i

2go

- 208

3~ m/z

I.U O

Z

186.

6e. I',-I LU

88

/~

Acoo.

A~ ~ - - _ _ _ 1

263

88

48.

O'[~

OOA~

551

> 300 x 10..

ZO

fl:: 8

376

A~

LJJ " >

160

160

B


188

146

.........L . ,,,., L4~_--.:-.L IBO

263 J

I

dd~ i~q:l 2BB

3~e

332 31~ [ ;1 _ ii

9

"3 7 6 .,.,

.,.

l

_

,

4ee

4is . , .

. . .

479 4~3 ( , . . . .

see

_

. . . . .

551 , .

.

.

.

see

m/z

Figure 9. GC-MS spectra of Man4GlcNAc after trifluoroacetolysis, NaBH4-reduction, periodate oxidation, NaBD4-reduction and peracetylation. (A) Branch connected to the 3position of the 3,6-disubstituted mannose residue. (B) Branch connected to the 6-position of the same disubstituted residue. Reprinted from: Anal. Biochem. 200 (1992) 58. Copyright 1995 Academic Press.

5.3. Mass

spectrometry

of O-linked

oligosaccharides

Oligosaccharides linked to Ser/Thr are released by alkaline borohydride as alditols, which is an advantage when structural analysis is carried out by periodate oxidation and mass spectrometry. To illustrate the use of this method on O-linked structures the following linear pentasaccharide alditol was chosen: Gall31-3GlcNAc]31-3Gall31-4GlcNAc[31-6GalNAc-ol Analysis of the intact compound by FAB-MS as permethylated derivative gave spectrum A in Figure 10. Primary sequence ions from the non-reducing terminal of m/z 464, m/z 668 and m/z 914 show a linear tetrasaccharide sequence. An alditol-containing ion of m/z 276 shows a monosubstituted GalNAc-ol and combined with the alditol-containing ions of m/z 521 and m/z 971 the linear sequence is confmned. The substitution positions of the GlcNAc residues can be

44 determined from the intensities of the secondary fragments. As previously discussed secondary fragments derived from a HexNAc residue are formed by preferential elimination from the 3position of this residue [2]. The primary sequence ion of m/z 464 gives a secondary of m/z 228 showing a Gall-3GlcNAc 1- sequence. A secondary ion of m/z 882 formed from m/z 914 by elimination of methanol shows that GlcNAc residue adjacent to GalNAc-ol is 4-substituted. No other linkage positions can be deduced from the spectrum. The monosaccharide composition is confirmed by an [M+I] + ion of m/z 1207. The FAB-mass spectrum of the oxidised compound as permethylated derivative is shown in spectrum B. The sequence ions from the non-reducing terminal of m/z 424 and m/z 228 show a Gall3GlcNAcl-sequence. The secondary fragment of m/z 228 determines the linkage position between the residues in agreement with data obtained from the intact compound (Spectrum A). The sequence ion of m/z 628 means addition of 204 m.u. to m/z 424 and consequently a periodate resistant residue which in this case is a 3-substituted Gal residue. The primary sequence ion of m/z 873 and a secondary of m/z 841 formed from the primary by elimination of methanol show a 4-substituted GlcNAc residue in accordance with the data obtained from the intact compound. From the ion of m/z 873 and an [M+I]+ ion of m/z 951 it can be concluded that the alditol has been shortened to ethylene glycol, which is a product of periodate oxidation of a 6-substituted GalNAc-ol. Supporting evidence for the sequence and linkage positions is furnished by the alditol-containing ions of m/z 305 and m/z 754. In most O-linked structures the GalNAc-Ser/Thr is 3,6-disubstituted. After treatment with alkaline borohydride the GalNAc-ol in the released oligosaccharide alditol is cleaved by periodate giving two products (Table 2). This is examplified by the following trisaccharide alditol: Neu5Ac(x2~ 6 GalNAc-ol Gal~l ~ 3 Analysis of the products by GC-MS after permethylation gave spectra shown in Figure 11. Spectrum A shows a periodate oxidised non-reducing terminal Gal determined by the cleavage ions of m/z 179 and m/z 104. Ions from the alditol part of m/z 189 and m/z 130 show an Nacetyl tetrosaminitol, which is a product of cleavage by periodate of the GalNAc-ol [27]. The spectrum represents the structural element Gall-3GalNAc-ol of the original trisaccharide alditol. The other part of the compound gave spectrum B. As previously discussed the oxidised sialic acid is recognised by the primary and secondary ions of m/z 289 and m/z 257, respectively. An alditol-containing ion of m/z 60 shows an ethylene glycol residue. The base peak of m/z 306 is a loss of a methoxycarbonyl radical from the moleuclar ion. These ions show that the GalNAc-ol is substituted in the 6-position by a sialic acid residue, establishing the structural element Neu5Ac2-6GalNAc-ol of the intact compound.

45

228

2 2 8 --- 4 6 4

668

, ,

uJ U Z r~ Z :::) <(: uJ > .J W -"

8 8 2 ---

914

70 68

971

521

276

58

]0

/196 I

,6, 432L

914

,,2 ~

521

4a

zlm

4,

[M+ll

6a

668

Bio

1207

8821

1

lia

~2r00

m/z

228 2 2 8 -,- 4 2 4 CH2OMe

80 w 0 Z Z

,..:

/,l MeODHC

78

o

~

CH2OMe

0

G0

< u,i > "I<:

so

r

]~

~O/~eO~o

~

~

0

CH~O ~

B

841 -- 873 CH,~O

!

~0

~O~?Me

~

J

~

Pur~n,J^

[,.,+23]

305

/

i.. LM+

" 24 196

951

"l~JlJ lJ/ I l l~,~ . 288

/ ........

O-CH2

~

754

48.

8

628

'

[ ' 368

4OO

754 ............ 500

512

liB_"; . . . . . . 600

663 695 7~?f;

/

[873 ~t;

I"~, ~,A . . . . . . . 788 880

1 I

I /

98

[ :~. . . . . . . 9OO

[89~

m/z

Figure 10. FAB-MS spectra of Gall 1-3GlcNAcl31-3Gall31-4GlcNAc[31-6GalNAc-ol. (A) Intact permethylated. (B) After periodate oxidation, NaBD4-reduction and permethylation. Reprinted from: Methods Enzymol. 193 (1990) 587. Copyright 1995 Academic Press.

46

104 -~2OM e

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)

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Figure 11. GC-MS spectra of Gal131-3[Neu5Aco~2-6]GalNAc-ol after periodate oxidation, NaBD4-reduction and permethylation. (A) Residue linked to the 3-position of the GalNAc-ol. (B) Residue linked to the 6-position of the GalNAc-ol. Reprinted from: Carbohydr. Res., 239 (1993) 35. Copyright 1995 Elsevier.

6. CONCLUSION In order to completely establish an oligosaccharide structure several structural parameters have to be determined e.g. linkage positions, sequence, identification of the monosaccharide building blocks and absolute- and anomeric configurations. One way to address some of these problems by mass spectrometry is to work on chemical modifications of the constituent monosaccharide residues in such a way that differences in mass lead to structural information, as an alternative to investment in costly instruments. Choice of ionisation technique is important, since the structural information depends on formation of fragment ions. El-MS and FAB-MS, in contrast to softer ionisation techniques, produce intense cleavage ions. Specific chemical pretreatments using periodate oxidation and trifluoroacetolysis have added another dimension to structural analysis of glycoconjugates by mass spectrometry, allowing determination of linkage positions in one single analysis. The chemical transformation used has to be specific and the yield of the reactions employed should be quantitative, otherwise wrong conclusions regarding the structure can be drawn from the spectra. Trifluoroacetolysis when used on high-mannose structures satisfies the demands. Periodate oxidation is a quantitative and specific reaction as determined by FAB-MS. However, some large and highly branched high-mannose structures sometimes give incomplete oxidation. This

47 problem could easily be resolved by further oxidation after reduction and complete oxidation was achieved.

7. R E F E R E N C E S 1

2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27

A-S. Angel, G. Gr6nberg, G. Nilsson, S. Str6mberg and B. Nilsson in Protein Glycosylation: Cellular, Biotechnological and Analytica Aspects. GBF Monographs. H.S. Conradt (ed.), VCH, New York, 15 (1990) 125. G. Gr/3nberg, P. Lipniunas, T. Lundgren, K. Erlansson, F. Lindh and B. Nilsson, Carbohydr. Res., 191 (1989) 261. H. Bj6mdal, C.G. Hellerqvist, B. Lindberg and S. Svensson, Angew. Chem. Int. Ed. Engl., 9 (1970) 610. I.J. Goldstein, G.W. Hay, B.A. Lewis and F. Smith, Methods Carbohydr. Chem., 5 (1965) 361. A-S. Angel, F. Lindh and B. Nilsson, Carbohydr. Res., 168 (1987) 15. A-S. Angel, P. Lipniunas, K. Erlansson and B. Nilsson, Carbohydr. Res., 221 (1991) 17. B. Nilsson and S. Svensson, Carbohydr. Res., 69 (1979) 292. B. Nilsson and S. Svensson, Carbohydr. Res., 72 (1979) 183. B. Lindberg, B. Nilsson, T. Norberg and S. Svensson, Acta Chem. Scand., B33 (1979) 230. B. Nilsson and S. Svensson, Carbohydr. Res., 65 (1978) 169. B. Nilsson and D. Zopf, Arch. Biochem. Biophys., 222 (1983) 628. P. Pfthlsson and B. Nilsson, Anal. Biochem., 168 (1988) 115. H. Wiegandt and G. Baschang, Z. Naturforsch., Teil B 20 (1965) 164. B. Nilsson and D. Zopf, Methods Enzymol., 83 (1982) 46. Y.-T. Li, Y. Ishikawa and S.-C. Li, Biochem. Biophys. Res. Commun., 149 (1987) 167. M. Ito and T. Yamagata, J. Biol. Chem., 261 (1986) 14278. A-S. Angel and B. Nilsson, Methods Enzymol., 193 (1990) 587. M.R. Hardy and R.R. Townsend, Proc. Natl. Acad. Sci. USA., 85 (1988) 3289. R.R. Townsend, M.R. Hardy, D.A. Cumming, J.P. Carver and B. Bendiak, Anal. Biochem., 182 (1989) 1. T. Mizuochi, K. Yonemasu, K. Yamashita and A. Kobata, J. Biol. Chem., 253 (1978) 7404. D.M. Carlson, J. Biol. Chem., 243 (1968) 616. Y.C. Lee and J.R. Scocca, J. Biol. Chem., 247 (1972) 5753. H. Krotkiewski, B. Nilsson and S. Svensson, Eur. J. Biochem., 184 (1989) 29. P. Lipniunas, A-S. Angel, K. Erlansson, F. Lind and B. Nilsson, Anal. Biochem., 200 (1992) 58. F. Maley, R.B. Trimble, A.L. Tarentino and T.H. Plummer Jr., Anal. Biochem., 180 (1989) 195. P. Lipniunas, G. Gr6nberg, H. Krotkiewski, A-S. Angel and B. Nilsson, Arch. Biochem. Biophys., 300 (1993) 335. H. Krotkiewski, E. Lisowska, G. Nilsson, G. Gr6nberg and B. Nilsson, Carbohydr. Res., 239 (1993) 35.

This Page Intentionally Left Blank

S.B. Petersen, B. Svensson, and S. Pedersen (Eds), CarbohydrateBioengineering 9 Elsevier Science B.V. All rights reserved.

49

Development of a novel enzyme based glucose sensor F. Spener, R. Steinkuhl, C. Dumschat, H. Hinkers, K. Cammann and M. Knoll Institut ftir Chemo- und Biosensorik Mtinster, Mendelstr.7, D-48149 Mtinster, Germany

Abstract

Glucose oxidase from Aspergillus niger is an extremely stable, FAD-dependent enzyme that has found wide application in electrochemical sensing of glucose in blood. Conventional sensor technology immobilises this enzyme in membranes deposited on top of the transducer. Here we report a new containment technology where the glucose oxidase is immobilised in gelatine or polyvinylalcohole and is deposited in the chip in pyramidal containments produced on silicon by anisotropic etching. This configuration of the sensor enhances the adhesion and stability of the membrane and has been applied to monitoring glucose in serum discontinuously as well as continuously in a flow-through system. Moreover we demonstrate the applicability of the containment technology to monitor glucose ex vivo with the help of a microdialysis system.

1. INTRODUCTION 1.1. Glucose oxidase

Glucose oxidase (I]-D-glucose: oxygen 1-oxidoreductase, EC 1.1.3.4) is a FAD-dependent enzyme that catalyzes the oxidation of ~-D-glucose by dioxygen to hydrogen peroxide and 8gluconolactone, which subsequently hydrolyzes spontaneously to gluconic acid. Apart from glucose monitoring in food, drinks and fermentation processes the most important application of this enzyme is in clinical diagnostics. Here it is used as a component of colorimetric diagnostic kits, of dry reagent test strips, and more recently of biosensors for the determination of glucose in blood, serum of plasma. Glucose oxidase (GOD) exhibits useful properties with respect to widespread application in biosensors because of the high specificity for glucose and an extremely high stability [1]. The enzyme has been isolated from various moulds, from red algae, citrus fruits, insects and bacteria. The most widly used enzyme in terms of research and commercialized products, however, is that from Aspergillus niger, a glycoprotein with a high-mannose type carbohydrate content of 10 to 16 % of its molecular mass. The carbohydrate component appears to be in form of a branched polysaccharide that partially surrounds the protein core [2]. Many of the enzyme's physical properties, such as high solubility in water and resistance to proteases for example, may be ascribed to its carbohydrate shell. Partial removal reduces the enzyme's stability.

50

Figure 1. Tertiary structure of FAD-containing glucose oxidase (monomer) from

Aspergillus niger [3]. The holo-enzyme is a homodimer, where each subunit contains the tightly bound (Kd = 1 x 101~ but not covalently attached coenzyme (FAD), and the reaction mechanism is essentially

H=O=,hydroquinones

Glucose " ~

S

GODo,-

= d ) ' -~ - ~

reduced mediators

,_ E ~

Fe(CN)=

~'~ ~

DCPIPH~

Glucose-GODo=

Glucono.~,,,..,,~

lactone

GOD~

DCPIP, low charge density Fe(CN)3, high charge density phenoxazines, ferrocenes,TTF, TCNQ 02, quinones

Figure 2. Natural and artifical substrates in redox reactions catalyzed by glucose oxidase from

Aspergillus niger.

51 dependent on this coenzyme. The recently unraveled tertiary structure of glucose oxidase (Fig. 1) from A. niger [3] shows that this enzyme allows electron acceptors other than dioxygen to reoxidize the reduced FAD in the same way. This is important for the application of the enzyme in the amperometric approach to glucose sensors, because the transfer of electrons between the active site of the reduced enzyme and an electrode takes place slowly or not at all. Transfer of electrons can be facilitated if a small electron acceptor is used as a mediator [ 1]. These compounds have the advantage that they allow amperometric biosensors to operate at relatively low potentials, and this can lead to a decrease in interference from electroactive compounds (Fig. 2).

1.2. Biosensors Given the extraordinary operational stability of glucose oxidase and the world-wide high demand for monitoring glucose in the blood of patients with type I diabetes (insulin-dependent diabetes mellitus) as a parameter for insulin injections that compensate the defect in insulin production by the pancreas, it is no surprise that the foremost application of biosensors to date is in the medical field. Initially biosensors for monitoring blood glucose have been used as analyzers in clinical laboratories. The US-company Yellow-Springs Instruments was the first to offer commercial glucose analyzers, based on the patent of L.C. Clark. Today the decentralized employment of point of care diagnostics in the doctor's office or in intensive care units and the bed-side measurements in hospitals are of greater interest (for example the multiparameter sensor of ISTAT Corp. Princeton, USA). Of utmost importance today, however, is self-testing by patients using miniaturized electrodes with immobilized GOD for blood glucose sensing. In this area 3 sensor systems are in different states of development. A big success already are the commercially available, disposable pen- or card-type sensors. The more provocative approach aims at continuous monitoring of blood glucose to attain normoglycemia and to avoid acute metabolic disturbances. With this in mind work on the development of implantable biosensors is going on for years. Due to the - in this case still limited - stability of the enzyme and still not optimal biocompatibility of all the sensor materials used, this concept has not yet been realized. A solution in the far future may be artificial organs, such as the pancreas, with an inbuild glucose sensor and insulin pump in nanotechnology. The more realistic concept for realization in the near future is a continuously working microdialysis system, were dialysates of the subcutaneuos tissue are channeled to a sensor for continuous glucose monitoring ex vivo [4]. Our approach to the glucose sensor is based on the GOD-catalyzed oxidation of glucose to gluconolactone and concomitant reduction of dioxigen to H202. The latter is electrochemically oxidized at a platinum anode. The anode is polarized at + 600 mV vs. an Ag/AgC1 reference electrode. Often the enzyme is immobilized in a membrane coveting the surface of the anode. The current depends linearly on the glucose concentration in the solution [5]. Until now most enzymatic biosensors have a membrane fixed on top of the transducer. This method often leads to malfunction of the sensor arising from problems like inadequate membrane adhesion and insufficient mechanical stability. Dipcoating procedures are difficult to perform in a reproducible way in order to obtain sensors with identical performances. Sensors with membranes that have separately been casted and mounted to an electrode can hardly be miniaturized and are therefore not suited for potential implantation.

52 In order to solve these problems we developed a new concept for membrane deposition [6, 7], the so called c o n t a i n m e n t s e n s o r s on the basis of micromechanically etched cavities in silicon substrates (<100> crystal orientation, 380 lam thick) with openings towards the analyte solution between 120 and 480 lam width, where the membrane is not located on but in the chip (Fig. 3). At the sides of the cavities platinum electrodes are deposited by means of semiconductor technology such as photolithography and physical vapour deposition. Here we demonstrate the application of this technology to the sensing of glucose in body fluids.

Pt Si SiO 2

Containment with immobilized enzyme

Figure 3. Schematic view of the containment sensor.

2. EXPERIMENTAL

2.1. Chip fabrication A silicon wafer (p-type, 3-inch, (100)-surface, 380 l.tm thick, optically polished on both sides) was oxidized in wet ambient at 1200 ~ for 210 min to get a 1.5 l.tm thick oxide layer. Spin-coating was performed sequentially with HMDS (hexamethyldisilazane) as adhesion promoter and photoresist (AZ 5214E/Hoechst) at 4000 rpm followed by a prebake at 90 ~ for 5 min. The resist layer had a thickness of 1.4 pm. It was exposed to about 80 mJ/cmz of mid UV radiation through a photomask to define the containments on the front side of the wafer. Development with AZ 524MIF/Hoechst was followed by a postbake at 120 ~ for 60 s. A protective coating was then applied to the back side of the wafer to withstand the following oxide etching process which was carried out with BOE (buffered oxide etchant) to get quadratic holes in the front side. Photoresist and protective layers where removed and silicon was etched anisotropically in 20 % KOH solution at 75 ~ for 6 h to create the containment holes. The silicon oxide was then stripped with hydrofluoric acid and a new 150 nm thick oxide layer was created thermally in dry oxygen at 1050 ~ In order to form platinum electrodes the front side of the wafer was first sputter deposited with a 1/am thick aluminum layer which acts as a sacrificial layer for reliable platinum patterning in a lift-off process. Using our newly developed electrospray-coating technique the aluminum layer was subsequently covered with a photoresist layer, again AZ 5214E from

53 Hoechst, which was now used in image 2 reversal mode. The prebake at 90 ~ for 5 min was sequentially followed by a 40 mJ/cm UV exposure through a photomask, a reversal bake at 120 ~ for 5 min, a 320 mJ/cm UV flood exposure, and development with AZ 524MIF. Then aluminum etching was performed with phosphoric acid etchant (PES 83.5-5.5-5.5/Merck) to get a negative mask for the following application of the platinum electrodes and conducting lines. A 200 nm thick platinum layer was then sputter deposited and patterned by removing the aluminum layer in a sodium hydroxide solution. Anodic bonding of a Pyrex | was performed at 500 ~ and 300 V for 60 min. Finally the chips were separated.

glass wafer

2.2. Enzyme immobilisation and membrane deposition The buffer (pH 7.0) was prepared by solving 8.1 g NaC1 (Merck), 0.272 g KH2PO 4 (Sigma), and 0.697 g K2HPO 4 (Sigma) in 1 1 deionized water. GOD (200 U/mg) was obtained from Fluka. 50 mg of acid photo gelatine (Filmfabrik Wolfen, Germany) were allowed to swell in 0.5 ml deionized water for 60 min at room temperature. Then 0.5 ml of the prepared buffer (pH 7.0) solution were added. After mixing the solution was kept at 35 ~ for 60 min, then different amounts of GOD-powder were added to 100 lal of the warm stirred gelatine solution in a small Eppendorf vessel. The vessel with the enzyme was placed in a chamber and the transducer chip was dipped into the enzyme-solution to cover the containment opening with it. Then the chamber was evacuated in order to fill the containment with the gelatine solution. After gelling the sensor surface was cleaned and the sensor was examined under a microscope to ensure a properly filled containment. Then the sensor was stored in buffer solution until the first measurement. Furthermore polyvinylalcohol functionalized with stilbene groups (PVA-SbQ) was used for enzyme immobilization [8]. 1600 U GOD were dissolved in 5 0 0 m g PVA-SbQ 500-88 solution. The solution was filled into the containment and then the PVA-SbQ was hardened by UV-irradiation. 2.3. Measurements The measurements were performed in a two electrode configuration. A chloridized silver wire was used as reference electrode. A voltage of 0.6 V was applied between the containment electrode and the Ag/AgC1 reference electrode (potentiostat Autolab PSTA 1 0 / 4 channel, Eco Chem). The sensors were tested in the buffer solution described above. Before starting calibration the sensors were polarized until the current was lower than 1 hA, which took about one hour. Known amounts of a highly concentrated glucose solution (Merck) were added to the stirred buffer solution to obtain the calibration graphs. When not in use the sensors were stored in buffer solution at 4 ~ The sensor was also probed in a flow system (fixed in a block of acrylic glass with a dead volume of approximatly 10 laD, equipped with a pump from Meredos and a valve from Knauer were used (the flow rate was 50 pl/min). The response time was obtained from the response curves in the flow system. The stability of the sensor was determined by immersing the polarised sensor together with the chloridized silver wire in stirred undiluted human serum (purchased from Sigma) and recording the current. The concentration changes in the undiluted serum were obtained by adding known amounts of high concentrated glucose solution.

54 3. RESULTS AND DISCUSSION The response curves and the corresponding calibration graphs of simultaneously measured containment sensors with different opening sizes (120, 250, 480 pm length of a side) toward the analyte solution are shown in Fig. 4. All sensors with gelatine based membranes responded fast and stable on changes in the glucose concentration. As expected the sensitivity of the sensors increased with increasing opening size. The detection limit was lower than 0.05 mM.

9

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10

Figure 4. Dependence of signal on the opening size (edge length) of the containment sensor toward the analyte solution. A, response curves; B, calibration curves. Glucose (0.139 - 9.12 mM) in buffer; 1600 U GOD/ml gelatine-solution.

In Fig. 5 it is demonstrated that the sensor is well suited for a flow system configuration. The sensor responded linearly and reproducibly in a concentration range relevant for continuous subcutaneous monitoring of glucose in a microdialysis system, if the microdialysis needle and flow rate are chosen properly. The response time (t90) in the flow system (Fig. 5) is 65 s for the change in concentration from 0.5 to 1.25 mM, and 104 s for the change from 1.25 to 0.5 mM.

55 60

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Figure 5. Response curve (A) and response time (B) of the containment sensor in the flow system. Glucose concentrations in buffer; 1600 U GOD/ml gelatine-solution; flow rate 50 pl/min.

160

.

,

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I

'

,

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Figure 6. Stability of the containment sensor during measurements of glucose in undiluted human serum. 2400 U GOD/ml gelatine-solution.

56 As demonstrated in Fig. 6, the sensor shows a stable signal in undiluted human serum over more than two days. This demonstrates the applicability of the sensor in undiluted body fluids. Because the linear range of the gelatine based membranes was not sufficient in undiluted probes, the photosensitive PVA-SbQ was chosen for the immobilization of GOD. The calibration plot indeed shows a wider linear range (Fig. 7). 100

I

. . . .

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20

30

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Figure 7. Response curve (A) and calibration curve (B) of the containment sensor with GOD immobilized in PVA-SbQ (1500 U/500 mg solution).

It is possible to store the sensors with GOD immobilized in PVA for more than 3 months in the refrigerator with a minute decline in sensor characteristics only. The influence of interfering substances was not investigated but a behavior similar to planar sensors using the same materials is expected. The results show that enzyme membrane deposition in silicon containments makes glucose sensors with good analytical response behaviour feasible. Linear range, in particular the normal physiological range of 3.9- 6.1 mM (70- 100 mg/dl) for glucose in blood and above in the diseased state, as well as lifetime and response times are reasonable well and acceptable for most practical applications, also for use in flow systems combined with microdialysis sampling. In the introduction we alluded to the lack of stable, implantable and long lasting glucose sensors [9, 10]. On the way toward this goal one immediate solution with application potential is the development of a miniaturized microdialysis system. Such a microdialysis system under development in our laboratory is shown in Fig. 8.

57

B

A gas-pressurized spnng

peffusion-solutio~ I

syringe

.

microcontainment-

f

microdia~/sis-ch~o

,,,

11

i

waste

C silicon t/on

Figure 8. Schematic view of the microdialysis system. A, Principle; B, lay-out of the microdialysis-chip; C, integrated microcontainment sensor.

The analytical response behaviour of these containment sensors is comparable with other sensor configurations which are fabricated in silicon planar technology, but offers the following advantages: - Containment sensors are very robust because the mechanically sensitive membrane is protected inside the chip against damage. Also adhesion problems with the membranes are minimized since they are attached inside the containment. Membrane patterning is not needed. - Encapsulation is no longer a problem since the main components of the sensor (membrane and metal electrodes) are located inside the containment. Encapsulation is further simplified -

58 due to the fact that the conducting leads and the sensitive sensor surfaces are located on opposite sides of the wafer. Thus, encapsulation can be performed on the top side of the whole wafer. - Containment sensors are ideally suited for integration into microsystems as the dead volume added to the system is minute. - The fabrication process is entirely compatible with mass-production technologies since it is a full wafer process. In mass production even the membrane deposition can be performed as full wafer process either under vacuum or with an automated dispense system. Only soldering the plug must be done after the chips have been separated. In conclusion, the containment concept allows the fabrication of glucose sensors with good analytical response behaviour and with a considerably improved technology.

4. REFERENCES

1 2 3

R. Wilson and A. P. F. Turner, Biosens. & Bioelectron., 7 (1992) 165. J.H. Pazur, K. Kleppe and A. Cepure, Arch. Biochem. Biophys., 111 (1965) 351. H.J. Hecht, H. M. Kalisz, J. Hendle, R. D. Schmid and D. Schomburg, J. Mol. Biol., 229 (1993) 153. 4 C. Meyerhoff, F. J. Mennel, F. Bischof, F. Sternberg and E. E. Pfeiffer, Horm. Metab. Res., 26 (1994) 538. 5 F. Scheller and F. Schubert, B iosensoren. Akademie-Verlag, Berlin, 1989. 6 M. Knoll, German Patent DE 4115414A1 (1991). 7 R. Steinkuhl, C. Sundermeier, C. Dumschat, K. Cammann and M. Knoll, German Patent application Nr. P 4337418.2 (1993). 8 R. Renneberg, K. Sonomoto, S. Katoh and A. Tanaka, Appl. Microbiol. Biotechnol., 28 (1988) 1. 9 G . S . Wilson, Y. Zhang, G. Reach, D. Moatti-Sirat, V. Poitout, D. R. Th6venot, F. Lemonnier and J.-C. Klein, Clin. Chem., 38 (1992) 1613. 10 S. J. Updike, M. C. Shults, R. K. Rhodes, B. J. Gilligan, J. O. Luebow and D. von Heimburg, ASAIO J., 40 (1994) 157.

S.B. Petersen, B. Svensson, and S. Pedersen (Eds), CarbohydrateBioengineering 9 Elsevier Science B.V. All rights reserved.

Carbohydrate

binding

at the active

site of Escherichia

59

coli

maltodextrin phosphorylase P. Drueckes, D. Palm and R. Schinzel Theodor-Boveri-Institut ffir Biowissenschaften der Universit~it Wtirzburg, Physiologische Chemie I, Am Hubland, D-97074 Wtirzburg, Germany

Abstract The oligosaccharide binding site of E. coli maltodextrin phosphorylase was characterised by a site-directed mutagenesis approach. Kinetic studies with oligosaccharides of different length imply that the oligosaccharide binding site consists of five subsites. Subsite five contributes about 14 kJ/mol to binding. Mutations of active site residues E67, E350 and H536 impair oligosaccharide binding significantly. Based on molecular modelling studies residues E350 and H536 appear to be important parts of subsite five, while residue E67 most likely contacts subsite two, which also seemed to be important for binding. Subsites three and four appeared to contribute only little to oligosaccharide binding energy.

1. INTRODUCTION Glycogen phosphorylases (EC 2.4.1.1) are key enzymes of the carbohydrate metabolism. They catalyse the phosphorolytic cleavage of t~-l,4-1inked glucose units to produce t~-Dglucose 1- phosphate: glucose, + P~ +--)glucosen_l + glucose 1-phosphate In muscle glucose 1-phosphate is further metabolised via glycolysis to provide energy, and in liver phosphorylase helps to maintain a constant blood glucose level via the action of glucose 6-phosphatase [4]. Therefore, suppression of glucose output from the liver may be achieved by inhibition of glycogen phosphorylase. Such inhibitors may be of use for therapy of the noninsulin dependent form of diabetes (NIDDM or Type II diabetes). One class of phosphorylase inhibitors consists of glucose analogs which stabilise the inactive T-form of the enzyme. Since there are only weak physiological inhibitors known, a variety of glucose compounds with better inhibitory properties were designed, synthesised and tested by L. Johnson, G. Fleet, N. Oikonomakos and coworkers [13, 23]. An alternate way to look for potent inhibitors other than glucose compounds would be to design analogs derived from the oligosaccharide substrate.

60 However, present knowledge about substrate binding sites of glucosyl residues in phosphorylases is still incomplete. From x-ray crystallography, molecular recognition and site directed mutagenesis studies the binding of glucose 1-phosphate and its derivatives in ground state and transition state is well characterised [20, 21, 3, 12]. Substantially less is known about productive binding of the polysaccharide awaiting degradation. So far no binding of oligosaccharides at the active site of rabbit muscle glycogen phosphorylase has been observed in crystals [18, 12], although there is at least gross structural information on a second carbohydrate binding site responsible for the attachment of carbohydrates to glycogen particles, the "glycogen storage site" [9, 12].

Synthesis

Degradation

Figure 1. Schematic drawing of the substrate binding site of maltodextrin phosphorylase. Subsites 5-2 are the primer subsites.

From kinetic studies with branched and linear oligosaccharides French and coworkers [7, 8] suggested a five glucose unit binding site, four subsites for the primer and one for the glucose moiety (Fig. 1). This subsite concept is comparable to that described for other oligosaccharide degrading enzymes [ 11, 16]. However, glycogen phosphorylases differ from those enzymes: First, in the reaction of phosphorylase a glucosyl residue is transferred to a phosphate group rather than to a water molecule. Consequently the exclusion of water from the active site is of essence for the phosphorylase reaction. Indeed, mutations of active site residues cause an increase of the remarkably low error rate, eg release of glucose rather than glucose 1phosphate [ 15]. Further, the phosphorolytic cleavage of oligosaccharides is freely reversible. At equilibrium (Kcq = Pi/Glc-1-P = 3.6 at pH 6.8) "synthesis" is favoured. In the cell the physiological role of phosphorylase is the energy conserving mobilisation of storage polysaccharides through phosphorolysis due to the relatively high phosphate concentration in the cell. The work presented here is aimed at mapping the oligosaccharide binding site by a combination of a kinetic studies with linear oligosaccharides of increasing length and sitedirected mutagenesis. The E. coli maltodextrin phosphorylase is used as a model system, since this enzyme binds short linear oligosaccharides better than glycogen phosphorylase. In addition, the bacterial enzyme lacks the glycogen storage site, which makes determination of kinetic parameters less complicated

61 2. SUBSTRATE SPECIFICITY OF MALTODEXTRIN PHOSPHORYLASE In the phosphorolysis mode the shortest oligosaccharide degraded by maltodextrin phosphorylase is maltopentaose (G5). Maltotetraose (G4) is degraded much less efficiently exhibiting an approximately 300-fold lowered specificity constant (Table 1). On the other hand no significant additional binding energy is provided by additional glucose units from oligosaccharides with more than five glucose residues (Table 1). A very slow degradation of maltotriose was detectable by thin layer chromatography but this activity was to low to be assayed quantitatively (Htilsmeyer and Palm, unpublished results). In phosphorolysis maltotetraose acts as an inhibitor with an inhibitor constant comparable to the Km values of the oligosaccharide substrate (Table 2). Maltotriose (G3), maltose (G2) and glucose (G) do not inhibit the phosphorylase reaction (Table 2). In the synthesis mode the phosphorylase reaction is efficiently primed in the presence of oligosaccharides with a minimum length of four glucose residues. [7]. Similar to degradation longer oligosaccharides do not have better primer properties. Maltotriose was found to be a markedly less efficient primer molecule. Comparable observations were made for potato phosphorylase [22]. No primer activity of maltose was detectable even at high enzyme concentration and at prolonged incubation times up to 48 h (Htilsmeyer and Palm, unpublished observations). In addition, maltotriose is an inhibitor of the synthesis reaction with a dissociation constant comparable to the Kin-value of the primer molecule maltotetraose (Table 2).

Table 1 Kinetic parameters of maltodextrin phosphorylase action on different oli~osaccharides Substrate Degradation Synthesis

G7 G6 G5 G4 G3

Km kcat (mM) (s -1)

kcat~m (s -1 mM -1)

Km keat (raM) (s -1)

kcat/Km (s "1 mM "1)

0.4 0.5 0.8 3.9 n.d.

45 44 38 0.13 n.d.

3.2 3.0 4.6 4.1 23

15 15 12,5 14,5 0.091

18 22 30 0.5 <0.01

48 44 58 60 2.1

A series of oligosaccharides ranging from G3 to G7 was labelled with dansylhydrazine at the reducing end. Labelled oligosaccharides (from G4 to G7) behave like physiological substrates in both directions. The fluorescence properties of the labelled substrates remained unchanged in presence of phosphorylase (Drueckes, unpublished results). Thus, the fluorophore of the substrates seems not to interact with the protein. This confirms the observation described above that additional glucose residues consecutive to G4 do not interact with the enzyme and that smaller oligosaccharides do not bind to the enzyme. From the kinetic and binding experiments the following model of oligosaccharide binding at the active site of phosphorylases emerges: Oligosaccharides which contain at least four glucose residues in synthesis or five residues in degradation, respectively, are required for effective

62 catalysis. Additional glucose residues do not contribute significantly to binding neither in synthesis nor in degradation. Therefore, the oligosaccharide binding site is composed of five

Table 2 Inhibitory effects of short oligosaccharides, maltose and glucose on the phosphorylase reaction Inhibitor Degradation Synthesis Ki (mM) Ki (mM) G4 0.4 G3 >100 4 G2 >250 >250 G >500 >500

consecutive subsites in the direction of degradation and four glucose binding subsites and one glucose 1-phosphate binding subsite in synthesis. The pronounced loss in catalytic competence between G4 and G3 or G5 and G4, respectively, points to a central role of subsite five in substrate binding both in the ground state and transition state. The contribution of this subsite to binding energy can be quantitated using AAGn = -RT In[ (kcat/Km)n_1//(kcat/Km)n] according to Horumi [11]. A free energy of binding of approximately 14 kJ/mol for subsite five was determined, while for subsite six and seven values of about 0,4 kJ/mol and 1 kJ/mol were determined. Since shorter oligosaccharides are no or only weak inhibitors, subsite five seems not sufficient to bind the oligosaccharide substrate and the other subsites must also contribute to oligosaccharide binding. If it is assumed that binding at two subsites is required for efficient oligosaccharide binding, all oligosaccharides which can occupy those sites should be substrates or at least inhibitors of the phosphorylase reaction. The failure of G3 to be an efficient inhibitor makes a contribution of subsite two to binding evident, since G3 cannot bind at sites two and five at the same time. Only G4 could span both binding sites thus making it an inhibitor molecule in the direction of degradation. If sites three or four would contribute significantly to binding, maltotriose should have inhibitory properties comparable to that of G4. Binding at subsites three and four seems not to provide binding energy or may even lower binding energy ("negative binding"). This is in accordance with observations of Giri and French [8] that subsite three can fit 1,6-glycosidic bonds. However, subsite four is specific for 1,4 linked glucose units. From these data no conclusions on the contribution of subsite one, the catalytic site, can be made.

3. FUNCTIONAL ROLE OF ACTIVE SITE RESIDUES IN OLIGOSACCHARIDE BINDING Since kinetic analysis of the oligosaccharide binding site is difficult due to the lack of suitable substrate analogs and the failure of earlier attempts to measure binding directly, a sitedirected mutagenesis approach was employed to get further information how carbohydrates bind at the active site. In an alanine scanning experiment first suggested by Cunnigham et al.

63 [5] a number of active site residues were changed to alanine (Ddickes et al., in preparation). Only those residues were selected for the mutagenesis experiments which meet at least two of three criteria: - Location at a channel with access to the active site which is most probably identical with the oligosaccharide binding site. This channel, which was proposed by Barford and coworkers [2], consists of an about 1,5 nm long channel from the cofactor binding site to the surface of the enzyme.

- Conservation in all known phosphorylase primary structures including enzymes of bacterial, plant and mammalian origins [ 14]. - Participation in oligosaccharide binding. Only amino acid residues with charged or hydrophilic amino acid side chains were changed, which are commonly involved in carbohydrate binding. In contrast to many other enzymes acting on oligosaccharides [19] no tryptophanes and only few aromatic amino acid side chains seem to be involved in carbohydrate binding by interacting with the plane of the sugar through nonpolar interactions. All mutant enzymes created so far have some properties in common: - They exhibited thermal stabilities comparable to that of the wildtype enzyme except for the E67A mutant enzyme, which is slightly more sensitive to heat. - There are no significant differences in the steady state parameters for maltopentaose and maltoheptaose (Fig. 2A) in the degradation mode or for maltotetraose and maltopentaose in the synthesis mode, respectively (Fig. 2B). This confirms the above mentioned observation that residues beyond the fourth glucose unit do not contribute significantly to binding of oligosaccharides.

- All mutations which influence carbohydrate binding affect binding of inorganic phosphate as well, but to a lower extent. This is in accordance with earlier observations that both substrates in the direction of degradation, inorganic phosphate and the oligosaccharide, do not bind independently [ 10]. 3.1. Effects of m u t a t i o n s on apparent binding According to their effects on apparent binding, the resulting mutant enzymes can be divided into three classes: Few mutant enzymes belonging to the first class showed no or only minor changes in the steady state parameters upon mutation although they are highly conserved in all phosphorylase sequences. In addition, one mutant enzyme, N112A, exhibits significantly increased thermal stability (Drueckes and Schinzel, unpublished results). The functional role of these side chains remains unclear. A second class is built of all mutations which caused only moderate changes of the apparent binding. The loss in apparent ground state binding energy (Fig. 2A) compared to the wildtype enzyme is about 4-6 kJ/mol for all mutants in this class. This corresponds to the loss of one

64 hydrogen bond between uncharged side chains [6, 20]. Indeed, all mutants belonging to this class are mutants of amino acids with hydrophilic but uncharged side chains: N307, Y578 and T346. In the case of the N258A/D259A/N260A mutant enzyme one of the asparagines rather than aspartic acid 259 might be engaged in binding of the oligosaccharide substrate in the ground state. Those three residues are part of a loop which in the inactive T-form of phosphorylase b blocks the putative substrate channel. This loop swings back upon allosteric activation of phosphorylase [2]. The moderate effects of this mutation on the non-allosteric bacterial phosphorylase shows that maltodextrin phosphorylase resembles the R-form of rabbit muscle phosphorylase. The Y578A and T346A mutants are of special interest since for these two mutants the binding energy in the ground state of the oligosaccharide in the synthesis mode is less affected than binding in the degradation mode (Fig. 2). This suggests that the oligosaccharide substrate may occupy partially different binding sites in synthesis and degradation, respectively. Although substitution of E350 by an alanine removes a charge, only moderate changes in apparent ground state binding are observed when compared to the E67A and H536L mutant enzymes. The third class consists of mutant enzymes which showed a more substantial decrease of apparent carbohydrate binding. Here the observed changes in free energy of oligosaccharide binding in the degradation mode are in the range expected for a hydrogen bond with a charged or uncharged residue, respectively (Fig. 2). This is the case for the E67A and H536L mutants, where charged amino acid side chains had been substituted by neutral ones. These data imply that both residues are of major importance in carbohydrate binding. However, in the direction of synthesis the values given (Fig. 2) represent only a lower limit since oligosaccharides are insoluble at high concentrations and thus Km values cannot be determined accurately. Therefore, conclusions about differential effects of these mutations on binding in synthesis and degradation cannot be drawn from the present data. 3.2. Effects of mutations on kcat/Km Changes in kcat/K m c a n be expressed in terms of the free energy of binding: AAG = - RT In[ (kcat~m)mut/(kcat/Km)wt]. In Fig. 3. the effects of mutations o n kcat/K m values are shown. As observed for the Km values, mutations of H536 and E67 causes the largest effects on k~a~Km. Due to the very high Km values kcat]tKm w a s determined directly at [S]<
65 activity. This suggests that a negative charge at this position is indispensable for a fully active enzyme (Drtickes and Schinzel, manuscript in preparation).

Figure 2. Effect of different mutations on Km values for oligosaccharides relative to the Km of the wildtype enzyme. The values for the H536L and E67A mutant give lower limits, since the determination of the apparent Km values was limited by the solubility of the oligosaccharides. (A) Degradation mode; (B) Synthesis mode. G4 = maltotetraose; G5 = maltopentaose; G7 = maltoheptaose.

The D308A mutant enzyme retained wildtype Km while activity is lowered 100-fold. Substituting T346 by alanine affects both binding in the ground state and to a larger extent binding in the transition state. Exchanging Y538 and N307 to alanine affects lqat only

66 moderately, the changes in AAG therefore reflect only the impaired binding in the ground state. Both residues seemed not be involved in transition state binding.

Figure 3. Changes in free energy of activation (calculated from kcat/K m values) of mutant maltodextrin phosphorylase compared with the wildtype enzyme. The values for the H536L and E67A mutant were determined directly at S<
3.3. Structural implications Unlike glucoamylases and or-amylases [17] the glycogen phosphorylases - although highly conserved from bacterial to mammalian enzymes- share no homology with other carbohydrate degrading enzymes. The binding site differs from binding sites in other enzymes acting on oligosaccharides in some respects, for example no tryptophan residues are located near the putative carbohydrate binding site in at the active site of glycogen phosphorylase. Up to now no structure of a phosphorylase - carbohydrate complex has been solved. However, a channel from the active site to the surface of the enzyme had been identified in the phosphorylase b structure [ 1] which is most probably the substrate channel. A maltopentaose molecule had been modelled into the phosphorylase b structure. However, this was only possible under the assumption that the oligosaccharide molecule is distorted mainly between the second and third glucose unit (L. Johnson, personal communication). Since almost all amino acid residues of the catalytic domain of the bacterial and mammalian phosphorylase are conserved [14], the assumption can be made that the 3-D structure of the active sites of both enzymes should be very similar. Therefore, the position of the amino acid residues which correspond to the mutated residues in the bacterial enzyme relative to the maltopentaose molecule was examined.

67 In this model H571 (corresponding to H536 in maltodextrin phosphorylase) and E382 (E350) are relatively close to glucose residue five. Tyr613 (Y578) appears to distant to make direct contacts to the sugar model. Hence the participation of this residue to substrate binding is most likely indirect or mediated by other side chains. In contrast, in the model only one side chain, E382 (E350), seems to be in the vicinity of glucose residues four. Binding at this subsite should contribute only little to binding both in the ground state and the transition state. Comparable to subsite four in the model only one amino acid, aspartic acid at position 339 (D308), is located near the glucose residue at subsite position three. However, the glucose residue at subsite two is in contact to all amino acids found to be involved in substrate binding in the bacterial maltodextrin phosphorylase (except for H571 and E382). From this observation it can be predicted, that subsite two contributes substantially to binding of the substrate and is critical to bind the oligosaccharide molecule in a distorted form. G5

~1376

-r

)-

PLP

Figure 4. Position of amino acid side chains (rabbit muscle numbering) found to be involved in oligosaccharide binding in maltodextrin phosphorylase to a maltopentaose molecule fitted into the structure of phosphorylase b (L. Johnson, personal communication).

68 If the kinetic and site directed mutagenesis studies are combined with the structural model of substrate binding (Fig. 4), a functional model of substrate binding in glycogen phosphorylases emerges: Subsites five and two seem to be of major importance for binding of the oligosaccharide in both the transition state and the ground state, while subsite three and four make no or only small contributions to binding. The different Km values observed for some mutant enzymes in the direction of degradation and synthesis point to a structural rearrangement of parts of the oligosaccharide binding energy site in catalysis.

4. ACKNOWLEDGEMENT We are grateful to L. Johnson, Oxford, for making the coordinates of the oligosaccharidephosphorylase b model available to us. This work was supported by a grant from the EC BIO2-CT94-3025.

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D. Barford, J.W. Schwalbe, N.G. Oikonomakos, K. Acharya, A. Hajdu, G. Papageorgiou, J.L. Martin, A. Knott, A. Vasella and L.N. Johnson, Biochemistry, 27 (1988) 6733. D. Barford and L.N. Johnson, Nature, 218 (1989) 233. S. Becker, D. Palm and R. Schinzel, J. Biol. Chem., 269 (1994) 2485. M. Browner and R.J. Fletterick, Trends Biochem. Sci., 17 (1992) 66. B.C. Cunnigham and J.A. Wells, Science, 244, (1989) 1081. A.R. Fersht, J.P. Shi, J. Knill-Jones, D.M. Lowe, A.J. Wilkinson, D.M. Blow, P. Brick, P. Carter, M. Waye and G. Winter, Nature, 314 (1985), 235. D. French and G.M. Wild, J.Amer.Chem.Soc., 75 (1953) 4490. N.Y. Giri and D. French, Arch. Biochem. Biophys., 145 (1970) 505. E.J. Goldsmith, S.R. Sprang, R. Hamlin, N.-H. Xoung and R.J. Fletterick, Science, 245 (1989) 528. D.J. Graves and J.H. Wang, The Enzymes, Vol. III, pp. 460-463, Academic Press, New York (1970). K. Hiromi, Biochem. Biophys. Res. Commun., 40 (1970) 1. L.N. Johnson, FASEB J., 6 (1992) 2274. J.L. Martin, K. Veluraja, L.N. Ross, L.N. Johnson, G.W.J. Flett, N.G. Ramsden, I. Bruce, M.G. Orchard, N.G. Oikonomakos, G. Papageorgiou, D.D. Leonidas and H.S. Tsitoura, Biochemistry, 31 (1991) 10101. C.B. Newgard, P.K. Hwang and R.J. Fletterick, Crit. Rev. Biochem. Mol. Biol., 24 (1989) 69. D. Palm, S. Becker and R. Schinzel, in Enzymes Dependent on Pyridoxalphosphate and other Carbonyl Compounds as Cofactors T. Fukui, H. Kagamiyama, K. Soda and H. Wada (eds.), pp. 377-385, Pergamon Press, Oxford (1991). M.R. Sierks, C. Ford, P.J. Reilly and B. Svensson, Prot. Eng., 2 (1989) 621. M.R. Sierks, C. Ford, P.J. Reilly and B. Svensson, Prot. Eng., 6 (1993) 75.

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S.R. Sprang, S.G. Withers, E. Goldsmith, R.J. Fletterick and N.B. Madsen, Science, 254 (1991) 1367. J.C. Spurlino G.-Y. Lu and F. Quiocho, J. Biol. Chem., 266 (1991) 5202. I.P. Street, C.R. Armstrong and S.G. Withers, Biochemistry, 25 (1986) 6021. I.P. Street, K. Rupiz and S.G. Withers, Biochemistry, 28 (1989) 1581. T. Suganuma, J.-I. Kitazone, K. Yoshinaga, S. Fujimoto and T. Nagahama, Carbohydrate Res., 217 (1991) 213. K.A. Watson, E.P. Mitchell, L.N. Johnson, J.C. Son, C.F. Bichard, M.G. Orchard, G.W. Fleet, N.G. Oikonomakos, D.D. Leonidas, M. Kontou and A. Papagoergioui, Biochemistry, 33 (1994) 5745.

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71

The chitinolytic system of Streptomyces olivaceoviridis H. Schrempf Department of Biology, University of Osnabriick, Barbarastral3e 11, D-49069 Osnabriick, Germany

Abstract

Streptomyces olivaceoviridis secretes several chitinases. The characterized 59 kDa exochitinase consists of a C-terminal catalytic domain, one central region, and one N-terminal chitin-binding part (12 kDa). The presence of the binding domain was shown to be a prerequisite for an efficient degradation of crystalline ~- and [~-chitins. In addition to the chitinolytic enzymes, S. olivaceoviridis produces a so far unique chitin-binding protein (CHB 1). Biochemical studies and immunofluorescence microscopy revealed that the CHB1 protein binds specifically only to c~-chitin from crab shell or fungi. The role of the novel lectinlike protein is at present being investigated.

1. INTRODUCTION Chitin, a polymer of N-acetyl-D-glucosamine, is highly abundant in nature, since it is present in the exoskeleton of insects, in mollusca, coelenterata, nematodes, protozoa and the cell walls of certain fungi. Naturally occurring chitins vary in the length of their chains which are stabilized by hydrogen bridges to a highly ordered crystalline structure. Chitin is frequently associated with proteins and may be stabilized by additional inorganic compounds. The natural annual production is judged to amount to 108 tons. Thus, apart from cellulose, it constitutes the second most abundant polysaccharide in nature [14]. Chitin can be hydrolyzed by enzymes produced by plants, fungi, and bacteria. Gram-positive streptomycetes are highly abundant in soil and known as important antibiotic producers. Though nearly all Streptomyces species have been shown to be chitinolytic, and chitin has been successfully used to enrich predominantly streptomycetes from soil [11], relatively few studies on chitinolytic enzymes have been performed. Streptomyces olivaceoviridis was identified as the most efficient degrader of crystalline chitin [ 1]. Recently, five chitinases (20.5 kDa, 30 kDa, 47 kDa, 70 kDa and 92 kDa) from S. olivaceoviridis have been purified to homogeneity [ 17]. Many extracellular hydrolases produced by bacteria consist of two or more domains. Most details are known about the catalytic and the binding domains of cellulases (CBDs). Proteins predicted from different bacterial chitinase genes showed in various cases an architecture of several domains, and comparisons of the deduced aminoacids allowed the identification of the

72 individual catalytic domains [23, 2]. With the help of biochemical and genetic data, chitinbinding domains could be identified within chitinases from Bacillus circulans [24]. We succeeded in analysing the catalytic and binding characteristics of an overproduced exochitinase and a novel lectin-like chitin-binding protein from S. olivaceoviridis as well as their corresponding chitin-inducible genes.

2. RESULTS AND DISCUSSION 2.1. The exochitinase and its gene Since S. olivaceoviridis degraded chitin efficiently, its DNA was used for shot-gun cloning experiments. In the presence of chitin, the S. lividans transformants containing the plasmid pCHIO 1 produced like S. olivaceoviridis a chitin-inducible exochitinase of 59 kDa which was proteolytically processed to a 47 kDa protein in the culture medium [2, 3]. Sequence analysis of a reading frame of 1794 base pairs, comparison of the deduced aminoacid sequence, and biochemical studies of the mature protein (59 kDa) and the proteolytically processed form (47 kDa) allowed the identification of one N-terminal chitinbinding domain, one central region with significant similarity to the type III module of fibronectin, and the C-terminal catalytic domain which spans from residues 261 to 597. This catalytic domain belongs to family 18 of glycosyl hydrolases [10] and four amino acids (Ser, Gly, Asp and Glu) were found to be completely invariable in the 12 aligned sequences. It is interesting to note that Asp and/or Glu residues have been discovered in the catalytic region of many glycosyl hydrolases including type-C lysozyme which also hydrolases chitin [22]. The deduced S. olivaceoviridis enzyme has highest overall amino acid identity with the B. circulans chitinase D [2]. The 59 kDa enzyme was shown to adhere very strongly to crystalline chitin and to hydrolyse native crab and fungal m-chitin more efficiently than the 47 kDa truncated form of the enzyme. In contrast, colloidal chitin and chitooligomers were degraded equally well by both forms of the enzymes. Neither standard physiological conditions nor elevated NaC1 and detergent concentrations allowed the removal of the bound Streptomyces exo-ChiO1 (59 kDa) enzyme; only high concentrations of urea and guanidine hydrochloride led to its release. Therefore it has to be assumed that the binding is irreversible under physiological conditions. These data reveal that a strong adhesion of the large form of the enzyme is a requisite for effective hydrolysis of the crystalline structure of chitin. In addition, it can be concluded that the N-terminal domain (12 kDa) of the 59 kDa enzyme is a chitin-binding domain [3]. By immunofluorescence, we could demonstrate that the 59 kDa chitinase mediates a very specific and strong binding to crystalline ~- and [3-chitins from various sources, but does not adhere to colloidal chitin. Since streptomycetes are dominant in soil and expected to encounter fungi in their natural habitat, it was not surprising that the 59 kDa enzyme interacts particularly efficiently with o~-chitin containing spores and phialides from fungi (i. e. Aspergillus proliferans and others). It hydrolyses the chitin within the fungal cell wall very well; thus protoplast-like structures and shortened mycelia are formed. We show for the first time that fungal chitin was hydrolysed considerably more efficiently than crab chitin which is usually used for the analysis of chitinase activities [3].

73

2.2. The lectin-like chitin-binding protein and its gene In the presence of chitin only, several chitinases including the exochitinase described above as well as an extracellular chitin-binding protein (CHB1) were produced by Streptomyces olivaceoviridis. The corresponding gene (chbl) was cloned on a multicopy vector in S. lividans. Like the exochi 1 gene, the chb 1 gene is only transcribed in the presence of chitin in its original host S. olivaceoviridis and the S. lividans transformant harbouring the chb 1 gene on a multicopy vector. Since the S. lividans transformant overproduced the protein only in the presence of chitin, it was rapidly purified to homogeneity [20]. Using immunofluo-rescence microscopy and biochemical analyses, a very specific binding of the overproduced CHB 1 to o~chitin was demonstrated. Native c~-chitin from crab shells consists of N-acetylglucosamine polymer chains arranged in an antiparallel way. Weaker adsorption of CHB 1 was found with colloidal chitin; this derives from native crab shell (x-chitin after treatment with acids, and its structure is probably predominantly amorphous. The CHB1 protein does not bind to carboxymethylchitin and to g-chitin, nor to any other type of cellulose. Since the CHB1 protein strongly interacted with o~-chitin and additional spectroscopical studies suggested a possible involvement of tryptophane residues with the crystalline substrate, we looked for a region within CHB1 harbouring several of these aminoacids. Four tryptophanes were shown to be deduced in the N-terminal part of the CHB 1 protein. By manual alignment of the relative positions of aromatic aminoacids within CHB1 and other glycohydrolases, four tryptophane residues were found in the cellulose-binding domains (CBDs) of several bacterial cellulases [8, 12, 13, 15, 25, 26]. The CHB 1 protein contained within a region of about 100 aminoacids one cysteine, two asparagines, and one glycine. These aminoacids are also present within several cellulose-binding domains (CBDs), each of which consists of approximately 100 aminoacid residues. Thus, they are about 70 aminoacids shorter than the CHB1 protein. It is interesting that two tryptophane residues are present in lectins from plants, such as Vicia faba [21] and Dolichos biflorus [19]. Several conservatively substituted aminoacids show an analogous arrangement of the above-cited CBDs and CHB 1. Lectins from the seeds of several plants (Graminaceae, Solanaceae and some Leguminosae) had been found to bind to N-acetylglucosamine, oligomers of this sugar and chitin [6]. One of the best-characterized chitin-binding lectins of plant origin is the wheat germ agglutinin (WGA). Within lectins, 30 - 43 aminoacids which contain various conserved cysteines and glycines (see also below) have been identified [4]. Analyses of deleted genes and their products from the B. circulans chitinases A1 and D resulted in the identification of chitin-binding domains of 52 to 55 aminoacids [24]. The exochitinase from S. olivaceoviridis [2,3] was shown to be proteolytically processed to a catalytically active part of the enzyme and a chitin-binding part of 131 aminoacids (see above). Up to now, the binding capacities of the domains from the above-mentioned chitinases have been tested with a-chitin, but neither with any other forms of chitin nor with different cellulose types. Despite our alignments, we could not identify similar aminoacid arrangements among the binding domains of the above chitinases and CHB 1. Further studies are necessary in order to test whether the various above-mentioned binding regions possibly mediate the adsorption to regions with a varying degree of crystallinity of the chitin and (or) if they differ in their interaction with regions which are largely or only to a small extent exposed to the surface of chitin. Although the sizes of the binding domains of various cellulases and chitinases of bacterial or eucaryotic origin range from 35 to about 150 residues, their g-structure can in

74 general be predicted (like for CHB 1, data not shown), and they seem to mediate the binding by exposed aromatic residues such as tryptophanes and/or tyrosines. More extensive studies are required to elucidate the structure of proteins interacting solely with a-chitin, in comparison with those binding to various types of cellulose and chitin. Therefore, truncated and mutated forms of the proteins are being investigated at present. Recent data give reason to believe that the cellulose-binding domains from cellulases are involved in the non-hydrolytic disruption of high-molecular weight cellulose. The precise action is not yet known, but it was proposed that the CBD protein binds to and penetrates the cellulose fibre at surface discontinuities and sluffs off cellulose fragments non-covalently associated with fibre but bound to underlying microfibrils [5]. Therefore the interaction of CHB 1 with chitin may assist the hydrolytic attack by enzymes and is being investigated at present. CHB 1 does not possess any antifungal activity. At present we are testing if it mediates the interaction of Streptomyces and chitin-containing fungi. It will also be interesting to analyse if the production of the specific chitin-binding protein is typical for S. olivaceoviridis, which had been isolated in the course of a screening programme [1] as an extremely efficient chitin degrader, or if other Streptomyces species produce binding proteins, too. Up to now, Calcofluor and Congo red [18] have been used to quickly identify polysaccharides within different organisms. However, the dyes are not very specific as they interact with various types of chitin, cellulose, and other polysaccharides. X-ray diffraction spectra have been necessary to distinguish different types of chitin. Since CHB1 binds specifically to a-chitin, it could also be used for the rapid identification of this chitin type. In addition, fluorescein-labelled CHB 1 could serve as a tool to study the synthesis of o~-chitin as well as the dynamics of its crystallization during the development of various organisms. Recent studies have revealed that cellulose-binding domains possess a large potential to purify various proteins. Thus a gene fragment encoding a CBD was fused to several genes encoding proteins of interest [16, 7]. Using similar constructs with the chbl gene, the adsorption of corresponding gene products to chitin could serve as a tool for the rapid enrichment of proteins which are difficult to isolate. Moreover, various proteins could be applied in their chitin-immobilized form.

3. ACKNOWLEDGEMENTS We thank M. Lemme for her help with the writing of the manuscript. The studies were supported by the Deutsche Forschungsgemeinschaft Schr 203/3-2.

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12 13 14 15 16 17 18 19 20 21 22 23 24 25 26

N. Din, N.R. Gilkes, B. Tekant, R.C. Miller Jr, R.A.J. Warren and D.G. Kilbum, Bio/Technology, 9 (1991) 1096. I.J. Goldstein and R.D. Poretz. In I.E. Liener, N. Sharon and I.J. Goldstein (eds.), The Lectins: Properties, Functions, and Applications in Biology and Medicine, Academic Press, pp. 35-250, 1986. J.M. Greenwood, E. Ong, N.R. Gilkes, R.A.J. Warren, R.C. Miller Jr and D.G. Kilbum, Prot. Eng., 5 (1992) 361. J. Hall and H.J. Gilbert, Mol. Gen. Genet., 213 (1988) 112. J. Hall, G.P. Hazlewood, N.S. Huskisson, A.J. Durrant and H.J. Gilbert, Mol. Microbiol., 3 (1989) 1211. B. Henrissat, Biochem. J., 280 (1991) 309. H.J. Kutzner, In M.P. Starr, H. Stolp, H.G. Trtiper, A. Balows and H. Schlegel (eds.), The Prokaryotes: A Handbook on Habitats, Isolation and Identification of Bacteria, Springer-Verlag, Berlin, Germany, pp. 2028-2090, 1981. A. Meinke, C. Braun, N.R. Gilkes, D.G. Kilburn, R.C. Miller Jr. and R.A.J. Warren, J. Bacteriol., 173 (1991) 308. A. Meinke, N.R. Gilkes, D.G. Kilburn, R.C. Miller Jr. and R.A.J. Warren, J. Bacteriol., 175 (1993) 1910. R.A.A. Muzarelli, Chitin, Pergamon Press, 1977. G.P. O'Neil, S.H. Goh, R.A.J. Warren, D.G. Kilburn and R.C. Miller Jr., Gene, 44 (1986) 325. E. Ong, N.R. Gilkes, R.C. Miller Jr., R.A.J. Warren and D.G. Kilburn, Biotechnol. Bioeng., 42 (1993) 401. A. Romaguera, U. Menge, R. Breves and H. Diekmann, J. Bacteriol., 174 (1992) 3450. C. Roncero and A. Dur~in, J. Bacteriol., 163 (1985) 1180. D.J. Schnell, D.C. Alexander, B.G. Williams and M.E. Etzler, Eur. J. Biochem., 167 (1987) 227. J. Schnellmann, A. Zeltins, H. Blaak and H. Schrempf, Mol. Microbiol., 13 (1994) 807. N. Sharon and H. Lis, Science, 246 (1989) 227. M. Sinnott, Chem. Rev., 90 (1990) 1171. T. Watanabe, W. Oyanagi, K. Suzuki, K. Ohnishi and H. Tanaka, J. Bacteriol., 174 (1992) 408. T. Watanabe, K. Kobori, T. Yamada, Y. Ho; M. Uchida and H. Tanaka, In R.A.A. Muzzarelli (ed.), Chitin Enzymology, Eur. Chitin Soc., Ancona, Italy, pp. 329-337, 1993. R.W.K. Wong, B. Gerhard, Z.M. Guo, D.G. Kilbum, R.A.J. Warren and R.C. Miller Jr., Gene, 44 (1986) 315. M.D. Yablonsky, T. Bartley, K.O. Elliston, S.K. Kahrs, Z.P. Shalita and D.E. Eveleigh, In J.-P. Aubert, P. B6guin and J. Millet (eds), Biochemistry and Genetics of Cellulose Degradation. FEMS Symposium 43, London, Academic Press, pp. 249-266, 1988.

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S.B. Petersen, B. Svensson, and S. Pedersen (Eds), Carbohydrate Bioengineering 9 Elsevier Science B.V. All rights reserved.

77

Properties and production of the [3-glycosidase from the thermophilic Archaeon Sulfolobus solfataficus expressed in mesophilic hosts M. Moracci a, L. Capalbo b, M. De Rosa b, R. La Montagna ~, A. Morana b, R. Nucci ~, M. Ciaramella ~ and M. Rossi a alnstitute of Protein Biochemistry and Enzymology, CNR, Via Marconi 10, 80125 Naples, Italy blstituto di Biochimica delle Macromolecole, II University of Naples, Via Costantinopoli 16, 80138 Naples, Italy

Abstract

The [~-glycosidase from the thermophilc Archaeon Sulfolobus solfataricus (Singly) shows broad substrate specificity, thermophilicity and high resistance to heat and other protein denaturants. It is an interesting model for elucidating the structure-function relashionships in glycosyl-hydrolases as well as the molecular determinants of thermophilicity and thermostability. In this paper we describe different strategies to obtain and optimize heterologous expression of the S[3gly in mesophilic hosts and down-stream processing, which are the pre-requisites for large scale production of the enzyme. We also describe the properties of the expressed enzyme and those of protein chimeras obtained by fusing the S~gly to the glutathione-S-transferase (GST); such fusions are useful tools to investigate on the properties of the enzyme and to easily purify mutants impaired in their activity or stability.

1. I N T R O D U C T I O N Among enzymes involved in carbohydrate metabolism, attention has been recently focused on [3-glycosidases from thermophilic microorganisms, enzymes showing highly thermostable [3glucosidase and [3-galactosidase activities. We have previously reported studies on the thermostable ]3-glycosidase (S[3gly) from the extreme thermoacidophilic Archaeon Sulfolobus solfataricus (for a review, see 1). The enzyme is thermophilic - the activity variation with temperature shows increase up to 90 ~ and thermostable - the half life at 75 ~ is 24 hours; it is stable to different protein denaturants and proteases, and moreover it is activated by heat and some protein denaturants. The enzyme has a broad substrate specificity: it hydrolizes [3-D-gluco-, fuco-, and galactosides, and a large number of [3-1inked glycoside dimers and oligomers, linked 131-3, [31-4, and [31-6. It has

78 noticeable exo-cellobiase activity on oligosaccharides with up to 5 glucose residues. Furthermore, it is also able to promote transglycosylation reactions [2,3]. The SI]gly encoding gene, called lacS, has been cloned and expressed in mesophilic hosts [4-6; see below]. We have analyzed the transcriptional organization of the lacS region, we have identified the transcription start site and the presumptive promoter sequences [7]. The enzyme has been crystallized [8] and the resolution of its structure is in progress. The SI3gly belongs to the Glycosyl-hydrolase family 1, including enzymes with 13glycosidase, 13-galactosidase and phosho-13-galactosidase activity, widely distributed among the three living Domains (Bacteria, Eukarya and Archaea). Enzymes of this family share significant sequence similarity with enzymes of cellulase family A, including microbial enzymes with 13-1,4 glucanase activity, which are involved in the degradation of cellulose. Enzymes of both families catalyze hydrolysis of 13-glycosidic bonds, possibly by the same mechanism of general acid catalysis [9]. For its particular features the SI3gly is an useful tool in all circumstances in which wide substrate specificity coupled with resistance to denaturation is required. For instance, its ability to hydrolyze oligosaccharides allows its utilization coupled with mesophilic or thermophilic cellulases in the conversion at high temperature of cellulose to glucose. We are interested in the identification and definition of the functional sites of the SI3gly involved in its different properties (substrate specificity, exo-cellobiase and transglycosylation activities), and to the structural deteminants of thermostability and thermophilicity; such results could also open the possibility of rationally engineering the enzyme to achieve better applicative performance. In this paper we describe different strategies for production, purification and downstream processing of the heterologously expressed SI3gly.

2. HETEROLOGOUS EXPRESSION The development of an innovative enzymology based on enzymes from thermophiles is currently hampered by difficulties in producing thermophilc biomass. The extreme growth conditions of hyperthermophiles and the slow growth rate with consequent low biomass yields, make it difficult both to obtain amounts of enzyme needed for structure-function studies and to devise methodologies for industrial scale up of production. Cloning and overexpression of thermophile genes in mesophilic hosts is now a well-established methodology to overcome these problems. Thermophilic enzymes are inactive at the host optimal temperature and the cells may allow the accumulation of higher quantities of the expression product. Moreover, the intrinsic stability of the heterologous enzymes allows their selective purification from the host proteins. 2.1. Expression in Escherichia coli Expression of protein genes in E. coli is the essential prerequisite for easy production of product for enzymological and structural studies, as well as for manipulation of the genes by random and site-directed mutagenesis. We have over-expressed the SI3gly in E. coli and have purified it to homogeneity by using a heating step which was proved to be crucial: most host proteins were easily removed and two chromatographic steps employed for the purification of

79 the native enzyme were by-passed [6]. Subsequent conventional hydrophobic chromatography, ion-exchange chromatography and gel filtration gave a purified protein with the same specific activity as the native enzyme, but with a better final purification factor and yield. A typical purification gave more than 5 m g ~ culture of homogeneous enzyme. Since the enzyme produced in E. coli shows the same properties of the native one, it can be used as a substrate for mutagenesis studies; by sequence alignment among members of the Glycosyl-hydrolase family 1 several conserved residues can be identified which are potential targets for site-directed substitutions.

2.1. Expression in yeast For its features, the SI3gly has several applicative potentialities in food and pharmaceutical industries; the possibility of expressing high levels of the enzyme in a safe (GRAS) host such as Saccharomyces cerevisiae is particularly attractive. We have previously obtained expression in yeast of the enzyme by using the vector pYE87 based on the classical URA3 selection [5]. However, when this expression system was scaled up to production volumes, it turned out that the amount of enzyme produced was significantly lower than expected, due to plasmid instability. To overcome this problem we have used an autoselection system based on the FBA1 gene, coding for the glycolitic enzyme fructose 1,6 bisphosphate aldolase, that is essential in any growth condition. In this system, the host strain harbors the chromosomal copy of the FBA1 gene stably disrupted [10] and contains a functional copy of the same gene on a plasmid. Since only cells retaining the vector are viable, the plasmid is completely stable after 50 generation in rich media [ 11]. We have constructed an autoselection-expression vector (pYG5) containing a functional copy of the FBA1 gene and the S. solfataricus lacS gene, coding for the SI3gly, under the control of the galactose-inducible UAScAL. Yeast cultures transformed with pYG5 or with pYE87 were subjected to three rounds of growth in non selective medium; with conventional selection plasmids, this experimental scheme favours high rate of plasmid loss (see below). In these conditions the two strains show very similar growth curve and reach the same maximal optical density (Fig.l), showing that the autoselection system does not result in growth disavantage. 12 10 9 pYE87 i pYG5

OF 0

. 20

40

i i 60 80 100 120 140 time

Figure 1. Growth curves of yeast cultures transformed with pYE87 and pYG5 vectors.

80 Plasmid stability was measured in cultures grown in non-selective conditions for 19 generations (Table 1). With plasmid pYE87, based on conventional selection, the fraction of plasmid-containing cells drops to 30%, whereas the autoselection vector pYG5 is present in 100% of the cells, confirming that this autoselection system is very stringent and that there are no plasmidless cells in population. The possibility of scaling-up of this expression system to production volumes will be evaluated.

Table 1 Stability of pYE87 and pYG5 in non-selective medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . gen_e_r___ati_ons......... 3 19 pYE87 70 % 30 % pYG5 97 % 98 %

3. D O W N S T R E A M PROCESSING

We have developed a down-stream processing methodology for the purification of the enzyme expressed in yeast, taking advantage of the unusual stability of the S]3gly to heat and proteases. This methodology involves two key steps: cell autolysis and extracts thermal precipitation. A pilot fermentation experiment was carried out with a S. cerevisiae recombinant strain expressing the Sl3gly in a 100 liter scale. Cells were collected, subjected to one cycle of freeze-thawing, and then to autolysis. During this process, the enzyme was released into the medium. Interestingly, the time-course of the protein released showed that, whereas total protein concentration decreases with time, presumably due to digestion by endogenous proteases released during the autolysis, the [3-glycosidase activity increased continously with time up to a maximum recovery after 72 hours, testifying once more the exceptional stability of the enzyme. The enzymatic yield,_in terms of units per liter of cell colture, is about 35-fold higher in the recombinant yeast autolysate than that obtained from the S. solfataricus homogenate. The enzyme was further purified by heating the autolysate for 20 min at 80 ~ and discarding the precipitate by centrifugation. The yield of the whole process (units/L cell culture) obtained with this new purification procedure increased up to 56-fold. This methodology for the recovery of a thermophilic enzyme from yeast is suitable for industrial processes, being inexpensive and acceptable for the environment.

4. GST-S]]gly FUSIONS: EXPRESSION AND PURIFICATION We have constructed fusions of the S~3gly to the glutathione-S-transferase (GST) from Schistosoma japonicum and we have obtained their expression in E. coli; such fusions allow at the same time one-step purification of the chimeric enzyme by affinity chromatography on glutathione, and assay of GST activity to follow purification. Therefore they offer an

81 alternative to classical methods for SI3gly purification, and are an essential tool for production of mutant enzyme molecules impaired in activity and/or stability. We made a protein chimera by cloning the lac S gene in the BamHI site of vector pGEX-2T [12], obtaining vector pGexgly. The resulting product contains at the N-terminus the 26 kDa GST polypeptide fused to the 56 kDa SI3gly protein. The sequence at the junction of the two polypeptides was as follows: -Leu-Val-Pro-Arg-Gly-Ser-Met-Tyr-Ser-Phe-Pro-Asnwhere the ftrst six lac S amino acids are in bold. After transformation of JM105 E. coli cells with pGexgly, the expression of the chimeric protein was obtained by induction with IPTG and, after over-night growth, abundant GST expression could be detected in crude homogenate at 37 ~ following the standard GST assay [13]. The extract was loaded on a Gluthatione Sepharose 4B column; no GST activity was released from the column after extensive washing and the fusion was eluted after injection of glutathione buffer. The sample obtained showed both GST and 13-galactosidase activity at 37 and 75 ~ respectively, and an high degree of purity. When the sample was analysed by SDSPolyacrylamide Gel Electrophoresis (SDS-PAGE) shown in Fig. 2, a two band pattern was found (lane 2): a 90 kDa band corresponding to the denatured chimera monomer, and a high molecular weight band which corresponds to the incompletely denatured, oligomeric form of the protein, which surprisingly is still active, as demonstrated by 13-galactosidase activity staining (lane 9). A similar pattern (but with bands of reduced size due to the absence of the GST portion) could be found for wild-type S[3gly analysed in the same conditions (lanes 7 and 8). These results demonstrate that the fusion of GST to Sl3gly has little effect on the overall architecture of the latter molecule, poorly affecting its functional folding, oligomerisation and exceptional thermal stability (fusions and wild-type protein samples, before loading on SDSPAGE, were incubated at 100 ~ in 0.1% SDS for 5 min.). Nevertheless, the ]3-glycosidase specific activity of the fusion was as much as 25% less than that of the wild-type enzyme at the same purification step, indicating that the catalytic efficiency and/or specifity were partially affected. In order to separate the GST portion from the fusion and recover the free SI3gly, samples obtained from the affinity chromatography were subjected to cleavage with thrombin protease. In fact, the fusion included a thrombin recognition site (bold residues): -Leu-VaI-Pro-Arg $ Gly-Ser-Met-Tyr-Ser-Phe-Pro-AsnThe arrow shows the bond hydrolysed by thrombin: as a consequence, the S]3gly produced by this digestion would show two additional amino acids at the N-terminus. Unfortunately the proteolytic cleavage resulted inefficient, even at very high thrombin/fusion ratio, probably because the thrombin recognition site is hidden into the chimera structure (Fig. 2, lanes 2 to 6). Indeed the efficiency of cutting was partially enhanced when the protein was pre-incubated at 55 ~ and then subjected to thrombin hydrolysis: in these conditions the GST portion is partially denaturated and presumably exposes the thrombin site. However, this treatment produced incomplete and aspecific protein cleavage (data not shown).

82

Figure 2. 10% SDS-PAGE of GST-SI3gly fusions after treatment with thrombin protease. Left panel: Coomassie staining. Lane 1, Molecular Weight Markers: Phosphorylase b 94,000 Da; Bovine Serum Albumin 67,000 Da; Ovalbumine 43,000 Da; Carbonic anhydrase 30,000 Da; Soybean Trypsin Inhibitor 20,100 Da. Lane 2, GST-SI3gly fusion incubated o.n. at 25 ~ Lanes 3-6, GST-SI3gly fusion incubated at 25 ~ in the presence of thrombin protease (10 units/mg of chimeric protein) for 1, 2, 4 hrs and o.n., respectively. Lane 7, wild-type SI3gly. Right panel: 13-galactosidase activity staining. Lane 8, wild-type SI3gly. Lane 9, GST-SI3gly fusion incubated o.n. at 25 ~ in the presence of thrombin protease (10 units/mg of chimeric protein).

In order to overcome this problem we prepared a new GST-SI3gly fusion, cloning the lacs gene in the BamHI site of vector pGEX-2TK, obtaining pGKgly. The resulting chimera protein, called GST-K-SI3gly, is similar to GST-SI3gly, but includes the recognition sequence for the catalytic subunit of cAMP-dependent protein kinase (PK) that is here underlined:

-Leu-Val-Pro-Arg $ Gly-Ser-Arg-Arg-Ala-Ser-Val-Met-Tyr-Ser-Phe-Pro-AsnWhen this new chimera was expressed and partially purified, it was efficiently and specifically cleaved by thrombin suggesting that the PK recognition sequence worked as 'spacer' between the two polypeptides, exposing the protease site (Fig. 3, lane 1). The 13glycosidase portion obtained in this way contained seven extra amino acids at its N-terminus. Nevertheless, this tail did not affect S~gly activity and thermal stability, conf'mrting that pGKgly vector can be used for the production of S[3gly mutants even severely affected in their properties.

83

Figure 3. 7% SDS-PAGE of GST-K-SI3gly fusions treated with thrombin protease, after Coomassie staining. Lane 1, GST-K-S[3gly fusion incubated o.n. at 25 ~ in the presence of thrombin protease (10 units/mg of chimeric protein). Lane 2, GST-K-S~gly fusion incubated o.n. at 25 ~ Lane 3, Molecular Weight Markers (see Fig. 2 legend). Lane 4, wild-type SI3gly.

5. CONCLUSIONS We have developed different strategies for optimization of the heterologous expression, improvement of the downstream processing, and analysis of the SI3gly, which offer us a versatile set of tools suitable for different applications. Our expression/purification system in yeast can be scaled up to production volumes, maintaining the foreign DNA stably in transformed cells, producing high biomass, high amount of the target product, consistency of yields in large volume fermentors, with low costs. Furthermore, the availability of GST-SI3gly fusions, besides interesting considerations on the structure-activity relationships of the wild-type enzyme, allows the purification of mutants designed to address questions about the enzyme activity, thermostability and thermophilicity.

6. ACKNOWLEDGEMENTS We are grateful to prof. E. Martegani for strains and suggestions. This work was partially supported by: CNR Target Project Biotechnology and Bioinstrumentation, and CNR P. F. Ingeneria Genetica.

5. REFERENCES

1 2

M. Moracci, M.Ciaramella, R. Nucci, L. H. Pearl, I. Sanderson, A. Trincone and M. Rossi, Biocatalysis, 11 (1994) 89. F.M. Pisani, R. Rella, C. Rozzo, C. A. Raia, R. Nucci, A. Gambacorta, M. De Rosa and M. Rossi, Eur. J. Biochem., 187 (1990) 321.

84 3 4. 5 6 7 8 9 10 11 12 13

R. Nucci, M. Moracci, C. Vaccaro, N. Vespa and M. Rossi, Biotechnol. Appl. Biochem., 17 (1993) 239. M.V. Cubellis, C. Rozzo, P. Montecucchi and M. Rossi, Gene, 94 (1990) 89. M. Moracci, A. LaVolpe, J. F. Pulitzer, M. Rossi and M. Ciaramella, J. Bacteriol., 174 (1992) 873. M. Moracci, R. Nucci, F. Febbraio, C. Vaccaro, N. Vespa, F. La Cara and M. Rossi, Enz. Microb.Technol., (1995) in press. A. Prisco, M. Moracci, M. Rossi and M. Ciaramella, J. Bacteriol., 177 (1995) 1614. L.H. Pearl, A. M. Hemmings, R. Nucci and M. Rossi, J. Mol. Biol., 229 (1993) 558. Henrissat, B., Biochem. J., 280 (1991) 309. C. Compagno, B. M. Ranzi and E. Martegani, FEBS Lett., 293 (1991) 97. C. Compagno, A. Tura, B. M. Ranzi and E. Martegani, Biotechnol. Progr., 9 (1993) 594. D.B. Smith and K. S. Johnson, Gene, 67 (1988) 31. B. Mannervik and U. H. Danielson, CRC Crit. Rev. Biochem., 23 (1988) 283.

S.B. Petersen, B. Svensson, and S. Pedersen (Eds), Carbohydrate Bioengineering 9 Elsevier Science B.V. All rights reserved.

85

Contribution of subsites to catalysis and specificity in the extended binding cleft of Bacillus 1,3-1,4-~3-D-glucan 4-glucanohydrolases Antoni Planas* and Caries Malet Laboratory of Biochemistry, Department of Organic Chemistry, CETS Institut Qufmic de Sarri?t, Universitat Ramon Llull, 08017 Barcelona, Spain

Abstract

Specific low molecular weight oligosaccharides having a chromophoric aglycon were synthesized for kinetic evaluation of the enzyme catalysis. A subsite mapping study showed that the active site cleft is composed of four subsites on the non-reducing end, and allowed to estimate the contribution of subsites - I I m -IV to transition state stabilization. In addition to a proper orientation of the scissile glycosidic bond of the substrate, inhibition kinetics with [3glucan- and cello-oligosaccharides suggest a binding preference of subsite -I for a 3-0substituted over a 4-O-substituted glucopyranose unit. Subsite +I, on the other hand, has a minor contribution to binding but the nature of the aglycon in the substrate largely affects catalysis.

1. INTRODUCTION 1,3-1,4-[3-D-glucan 4-glucanohydrolase (EC 3.2.1.73, abbreviated 1,3-1,4-[3-glucanase) catalyses the regio and stereospecific hydrolysis of [3-glucans containing mixed [3-1,3 and [3-1,4 glycosidic linkages as cereal 13-glucans and lichenan [1,2,3]. The trisaccharide 3-O-[3cellobiosyl-D-glucopyranose (1) and the tetrasaccharide 3-O-]3-cellotriosyl-D-glucopyranose (2) are the final hydrolysis products of barley [3-glucan, thus defining the enzymic specificity of cleavage on [3-1,4-glycosidic linkages in 3-O-substituted glucopyranose units.

I ~,,..OH FI H O ' 7 " .OH-~Z~o~O

(,,,OH

HO"~O~

OH

I (n=l)2 (n=2) The 1,3-1,4-[3-glucanases cloned and sequenced so far belong to two distinct enzyme families, plant and microbial enzymes, with no apparent sequence similarity and unrelated

86 three-dimensional structures. The Bacillus isozymes are the best characterized; they are highly homologous, with molecular masses of 25-30 kDa, active in a wide pH range, basic pI (8-9), and are quite thermostable proteins compared to the plant isozymes. Because of its importance in the food industry (brewing and animal feedstuff), the enzyme is a current target of protein engineering programs both to understand the enzyme function at the molecular level, and to redesign some enzyme properties for practical applications. Recent progress on the structure-function relationships in Bacillus 1,3-1,4-~-glucanases includes identification of the catalytic residues by site-directed mutagenesis [4,5] and chemical modification with active site-directed inhibitors [1], as well as determination of the tertiary structure by X-ray crystallography [6,7]. Bacillus licheniformis 1,3-1,4-~-glucanase is a retaining glycosidase [8] with Glu138 acting as the general acid catalyst and Glu134 as the catalytic nucleophile [5]. The three dimensional structure has been recently refined at 0.18 nm resolution by X-ray crystallography showing almost identical structure to that of the previously reported hybrid H(A16M) enzyme consisting of amino acid residues 1-16 from B. arnyloliquefaciens and the rest derived from B. macerans [6]. The protein adopts a jelly roll ~-sandwich fold with a carbohydrate-binding cleft located on the concave face of a ~-sheet formed of consecutive anfiparallel ~-strands. The catalytic residues Glu134, Asp136 and Glu138 are placed in the same ~-strand, and surrounded by a number of aromatic amino acid residues as it is commonly found in carbohydrate-binding proteins. The polysaccharide substrate may bind into the extended cleft which can be envisaged as formed of individual subsites each one accomodating a glucopyranose unit of the bound carbohydrate. The structure of a covalent enzyme-inhibitor complex between the hybrid H(A16M) 1,3-1,4-~-glucanase and 3,4-epoxybutyl ~-cellobioside [6] provides little information about the binding site cleft, since the cellobiose unit of the inhibitor only partially fills the cleft and it is covalently attached to the protein by an alkyl chain. Kinetic studies, on the other hand, have been limited to natural polysaccharide substrates, namely barley or oat ~-glucan, lichenan, and the reduced SIII pneumococcal polysaccharide (a regular polymer containing alternating 1,3- and 1,4-~-D-glucopyranosyl units [9]). These substrates undergo an extensive endo depolymerization with a large number of glycosidic bonds hydrolyzed per molecule yielding a complex mixture of oligosaccharides in a wide range of molecular weights at initial reaction times. Although they are currently used for analytical purposes (i.e. determination of I]-glucanase contents in cereals), the use of polysaccharide substrates is not appropriate for kinetic measurement in mechanistic studies of the enzyme action. We have recently prepared a family of low molecular weight I]-glucan oligosaccharides bearing a chromophofic aglycon as potential substrates for 1,3-1,4-~-glucanases. Here we describe their use in the enzymology of the Bacillus licheniformis enzyme aimed to evaluate the contribution of subsites in the extended binding site cleft to binding and catalysis.

87

2. RESULTS 2.1. Chromophoric substrates: 4-methylumbelliferyl 13-glucan oligosaccharides A set of low molecular weight 13-glucan oligosaccharides having a 4-methylumbelliferyl aglycon have been synthesized for kinetic evaluation of the enzyme catalysis [ 10]:

OH

r 0 3a

0

.OH'-]

?H

(,,OH

3b-e (n=0,3)

Criteria for designing the specific substrates were: a.-a basic substructure G3G-X according to the requirements of natural polysaccharide substrates (barley [3-glucan and lichenan) to be hydrolyzed by the enzyme, i.e. cleavage of a 13-1,4 glycosidic bond on a 3-O-substituted glucopyranose unit b.-single scissile glycosidic bond, so the enzymic reaction could be described by a simple kinetic model c.-release of a chromophoric aglycon upon enzymic hydrolysis for easy monitoring of the reaction course by continous UV or fluorescence spectrophotometry.

HPLC and NMR monitoring of the enzymic hydrolysis by B a c i l l u s l i c h e n i f o r m i s 1,3-1,4-13glucanase shows that 3a-e undergo a single cleavage with release of 4-methylumbelliferone, and that the reaction proceeds with net retention of the anomeric configuration. 2.2. Number of substites on the non-reducing end and their contribution to transition state stabilization Kinetics were performed at pH 7.3, 53 ~ conditions of maximum activity on the natural substrate barley 13-glucan. Initial velocities (<3% conversion) for 3a-e at different substrate concentrations were determined by monitoring the release of 4-methylumbelliferone at 365 nm (Ae = 5440 M-lcm-1). 3a is a very poor substrate, no saturation kinetics being observed up to 10 mM. The curve v o vs. IS] for 3b-e reaches a maximum and decreases at higher substrate concentration (Figure 1). The data were fitted to equation 1 that corresponds to an acompetitive substrate inhibition model. A clearly biphasic Hill plot, log[v/(V-v)] vs. log[S], (Figure 1, for 3d as an example) is consistent with this inhibition scheme, where a second molecule of substrate binds into the extended binding site cleft with formation of unproductive ternary complexes (ES2), producing a downward curvature of the Hill plot at high substrate concentration with the slope becoming negative (h =- 1). Kinetic parameters are summarized in Table 1.

88 2.0

k +l k2 E + S------~- E.S....___.~ E + p k-1 + S

1.6O~

1.5

1.2-

"~ 1.0

"-" x 0.8"

~E 0.0

E

,

b

h ~ = - I

"//

E.S

0.5-

vo =

-0.5" -1.0" -1.5

0.4-

kcat[Z~[Eo] K. +Is]+

w

-

!

-

|

-

(1)

Kt

i

-' .5-0.5 0.5 1.5 2.5 3.5 0.0-

In IS]

I

0

'

I

5

I

'

10 [S] (mM)

I

15

'

I

20

Figure 1. Steady-state kinetics for 3d. Inset: Hill plot

Table 1 Kinetic parameters of B.licheniformis 1,3-1,4-~-glucanase with substrates 3a-e. Substrate K M (mM) kcat (s "l) K 1 (mM) kcat/KM(M-Is -1) 3a

3b 3c 3d 3e

G-MUa G3G-MU G4G3G-MU G4G4G3G-MU G4G4G4G3G-MU

16.8+2.2 2.69+0.06 0.79+0.02 0.66+0.03

0.139+0.012 4.58+0.06 8.86+0.10 8.78+0.20

57.12+11.5 37.8+1.5 36.4+1.8 29.3+4.9

0.348+0.008 8.27+1.80 1700+60 11200+500 13300+900

a MU: 4-methylumbelliferyl. Conditions: citrate-phosphate buffer pH 7.3, 53~ 0.1 mM CaC12 [E]= 25-100nM, [S]" 3a 1-10 mM, 3b 1-51 mM, 3c 0.1-24 mM, 3d 0.1-22 mM, 3e 0.1-6 mM

The dependence of kcat and K M as a function of the degree-of-polymerization of the substrates approaches a plateau between the tetrasaccharide 3d and the pentasaccharide 3e. It inditates that the binding site cleft is composed of 4 subsites for glucopyranose units on the non-reducing end from the scissile glycosidic bond. This result was further analyzed by computer molecular modelling on the X-ray structure of the Bacillus licheniformis enzyme. Docking of a hexasaccharide into the binding cleft based on the 3D structure of the covalent protein-inhibitor complex between B.amyloliquefaciensB.macerans 1,3-1,4-[3-glucanase and 3,4-epoxybutyl I]-cellobioside [6] is consistent with a cleft containing 4 subsites, where a fifth glucopyranose unit on the non-reducing end is not conformationally restricted by the interaction with the protein in the productive complex, but rather it lays out the cleft and is solvent exposed.

89

Subsite mapping.

Binding to a multi-subsite binding site cleft of a depolymerase may lead to different positional isomers. The observed acompetitive substrate inhibition behaviour for compound 3b-e provides evidence for multiple binding modes, only one leading to a productive complex followed by specific hydrolysis at a single glycosidic bond. The general kinetics model accounts for all possible binary and ternary complexes, k +l,i,n ~_ E + Sn--. k -1,i,n

k2 ,r,n E'S i.n ' ' ' - ' ' - ~ +

E + Pn-1 +X

Sn

E'S i,n'Sj,n

where i and j indicate the binding mode (positional isomer), n is the length of the oligosaccharide, r denotes the productive complex, and E and Sn are the free enzyme and substrate. From steady-state considerations, the experimental kcat, K M and K I parameters relate to the microscopic constants of this model by

i :xr

(

KI -- ~lKr'n q" i,; ~-" 1/Ki'n

i :xr

)

l ~., ~_~1/Ki,nKi,j, n j

k 2,r,n

(kcatl=lk2,r I ~ KM ) ~. Kr )n

i

Kr, n and k2,r, n

being the microscopic Michaelis constant and the hydrolytic coeficient for the productive process, and Ki, n (ir r) and Ki,j,n the microscopic dissociation constants for the unproductive binary and ternary complexes. Only the second order rate constant (kcat/KM) is independent of the eventual formation of unproductive complexes. The subsite mapping model assumes that the free energy of binding at each subsite is an intrinsic constant, unaffected by binding or absence of binding at any other subsites [11], so - -Rr

2;

-

rAs.

h

AGh b being the contribution of a subsite h to the free energy of binding (AGnb) for a substrate of length n in the binding mode i. Because Kr, n and k2,r,n are not experimentally accessible, and assuming that the intrinsic binding energy can be used both for substrate binding and for lowering the activation free energy in catalysis, the contribution of each subsite to ligand

90 binding can not be calculated. Additional substrates to map the reducing end of the binding cleft are first required to complete the analysis and be able to estimate binding energies. However, the effect of occupying a subsite h on the second order rate constant kcat]K M is the contribution of that individual subsite to transition state stabilization, and can be expressed by the difference in transition state activation energy between two substrates differing in one glucopyranose unit according to AG*r,h=n+l = AG r,n+l ~ - AG ~ = - R T In r,n

(keat/KM )n+ 1 (kcat/KM) n

The values of AGh r are calculated from the data in Table 1 and summarized in Table 2.

Table 2 Contribution of subsites to transition state stabilization. Subsite Binding mode -I 3-O-Glcp -II 4-O-Glcp -III 4-O-Glcp -IV 4-O-Glcp -V

AGh* (kcal.mo1-1) -2.1+0.2 -3.5+0.2 -1.2+0.2 -0.11 +0.07

Figure 2. Subsites on the non-reducing end of the bound carbohydrate and their contribution to transition state stabilization.

Binding of 4-O-substituted glucopyranose units to subsites - I I - -IV have a stabilizing effect on the enzyme transition state complex with a larger contribution of subsite -III (Figure 2). A virtual subsite -V would contribute by less than 0.1 kcal.mo1-1, a value assigned to

91 unspecific interactions between the edge of the binding cleft and a fifth Glcp unit facing the bulk solvent in a loose conformation. Kinetics with substrates cannot give any information about subsite -I since it will be always occupied in the productive complex.

2.3. Contribution of substite -I to substrate specificity Being a heterodepolymerase, the contribution of each subsite varies depending on whether it is occupied by a 3-0- or 4-O-substituted glucopyranose unit. In addition to the proper orientation of the scissile glycosidic bond of the substrate, the strict substrate specificity shown by the enzyme suggests a preference for a 3-O-substituted glucopyranose in subsite -I. Inhibition kinetics with [3-glucan- (1, 2) and cello- (4, 5) oligosaccharides were analyzed to address the question of subsite selectivity. !

H

~

,.... OH

, I OH

F

~

~

c"OH

n

1 (n=l)

2 (n-2)

OH

ot4

4 (n=l)

"'OH

5 (n=2)

All of them behave as competitive inhibitors of B. licheniformis 1,3-1,4-]3-glucanase using the 4-methylumbelliferyl tetrasaccharide 3d as substrate. To avoid complex behaviour arising from substrate inhibition, the range of substrate concentration was chosen to be close to K M (0.33 mM < [S] < 1 mM, K M = 0.79 mM), far below K I (3d) = 36 mM (Table 1) under the same experimental conditions. Figure 3 shows the double reciprocal plot for 2 as an example. Inhibition constants are compiled in Table 3. 2

2.5-

1.5 2.0-

1

0.5 0

E 1.5-

[I] (mM)

i O

x o 1.00.50.0 -1.5

J '

I

-0.5

I

0.5

'

I

1.5

1/[S] (mM -1)

Figure 3. Competitive inhibition of 2 using 3d as substrate.

'

I

2.5

'

I

3.5

92 Table 3 Inhibition constants for the competitive inhibitors 1, 2, 4 and 5. Inhibitor K! (mM) 1 G4G3G 12.0+0.1 4 G4G4G 20.4_+0.3 2 G4G4G3G 2.28+0.04 5 G4G4G4G 5.60_+0.11 Conditions: citrate-phosphate buffer pH 7.3, 53~ 0.1 mM CaC12, [E]= 25nM, [S]=0.2-1 mM, [I]: 1 0-10 mM, 2 0-2 mM, 4 0-20 mM, 5 0-10 mM

The tetrasaccharides 2 and 5 are better inhibitors than the trisaccharides 1 and 4 as it may be expected for a glycosidase with an extended binding site. On the other hand, the [3-glucanoligosaccharides 1 and 2 having a [3-1,3 glycosidic bond have lower K! values than the corresponding cello-oligosaccharides 4 and 5. The change in free energy of binding associated with the structural modifications that relate any pair of inhibitors can then be calculated from the relationship A(AGb) = -RTln[(KI)x/(KI)y], where x and y refer to each compound. As shown in the thermodynamic cycle of Figure 4, the total free energy change in going from the worst inhibitor (G4G4G, 4) to the best one (G4G4G3G, 2) is independent of the path taken. However, the effect of each modification on binding depends on the pair of inhibitors considered. Thus, addition of a new glucopyranose unit to the non-reducing end of the ]3glucan-trisaccharide (1--)2, -1.08 kcal.mol ~ has a larger contribution than the same structural modification on the cello-trisaccharide (4--)5, -0.84 kcal.mol-1). Likewise, conversion of a ]3-1,4 to a 13-1,3 linkage has different effect for the trisaccharides (4--)1, -0.35 kcal.mol ~ than for the tetrasaccharides (5--)2,-0.58 kcal.mol-1). If the free energy of binding at each subsite is an intrinsic constant, unaffected by binding to any other subsite (subsite mapping model), this

G4G4G(4) ]

-0.84 "',,

-

~ [ G4G4G4G(5) ]

0 . 3 5 ,-'"

- 0.35 ss /

-

1.43

9

",,

G4G3G(1) ] Figure 4. A(AG b) in kcal.mo1-1 that relate the competitive inhibitors 1, 2, 4 and 5.

differential behaviour is interpreted as an evidence of different binding modes for each enzymeinhibitor complex.

93 Any microscopic interpretation should be handled with care since the measured K I values may be function of all possible equilibria between free enzyme and inhibitor, and a number of binary or even ternary complexes in different binding modes. A preferential binding mode, however, can be expected given the specificity of the enzyme. A binding model consistent with the above data is proposed and summarized in Figure 5.

(4) ~ ~ i ~ i ~ -IV

-III

-II

-I

AG_I (1___>4) ~ r

I

(-0.84)

9

~ 9

AG-I(I~3) - AG-Iv

js S

(-0.35)

oo oo

."

oS

I(

.r

~

(

5

-IV

-III

-II

(-0.58)

9149 9

) -I

I

AG-I(1---)3) - AG-I (1---~)

9

IIK

Figure 5. Proposed binding model describing the preferred binding modes. - O : 4-O-Glcp,-O" 3-O-Glcp. In parenthesis, A(AG b) in kcal.mo1-1

The 13-glucan-oligosaccharides 1 and 2 bind preferentially at subsites -I to -III or -IV, positioning the 13-1,3 glycosidic bond between subsites-I and-II as it may be expected from the substrate specificity shown by the enzyme. The A(AGb) value of-1.08 kcal-mo1-1 that relates 1 and 2 is thus the free energy of binding assignable to subsite -IV. This value is very close to that determined previously for AGe_IV (-1.2 + 0.2 kcal.mo1-1, see w suggesting that the contribution of this subsite to transition state stabilization (AGe_IV) is mainly due to stabilization of the enzyme-substrate complex. The cello-trisaccharide 4 will not bind as the ]3-glucan-trisaccharide 1. If subsite -I prefers a 3-O-substituted Glcp, and binding to subsite -IV is exoergonic, 4 will fill subsites -II to -IV. But, in going to the cello-tetrasaccharide 5, the additional glucopyranose may either lay out of the binding site cleft or fill subsite -I. Since K! for 5 is lower than for 4 (with an associated A(A G b) of-0.84 kcal.mol-1), the cello-tetrasaccharide will not bind to the same subsites -II to -IV because the fourth Glcp will have no contribution to binding, but rather it will fill subsite -I, resulting in a binding mode similar to that proposed for the 13-glucan-tetrasaccharide 2. Therefore, A(AG b) between 5 and 2 accounts for the preference of subsite -I for a 3-0substituted over a 4-O-substituted glucopyranose unit (-0.6 kcal.mol-1). From this model, the contribution of subsite -I to binding a 3-O-substituted glucopyranose unit of 13-glucan

94 oligosaccharides in the normal and preferential binding mode can be evaluated as being 1.4 kcal. mol- 1. The proposed model is tentative and requires further analysis, i.e. evaluation of the inhibitory capacity of the ct and [3 anomers, extend the analysis to larger oligosaccharides to test the consistency of the model, and determination of the 3D structure of some enzymeinhibitor complexes. 2.4. Subsite +I has a minor contribution to binding but a large contribution to catalysis The 4-methylumbelliferyl glycosides 3a-e reported as substrates (see w have an aromatic aglycon that fills subsite +I in the productive complex. To evaluate the effect of the leaving group in the hydrolytic reaction as well as the contribution of subsite +I to the enzyme specificity, compounds 6 and 7 were prepared and assayed with the B. licheniformis enzyme. Together with the 4-methylumbelliferyl trisaccharide 3c, these compounds share a common fragment ~-Glcp-(1-->4)-~-Glcp-(1-->3)-~-Glcp- that occupies subsites -I to -III in the productive complex (Figure 6). The methyl glycoside 6 is not hydrolyzed by the enzyme as determined by HPLC and NMR monitoring. On the other hand, the methyl glycoside 7 is a good substrate. Initial velocities were determined by monitoring the increase of reducing power due to the newly-formed reducing ends upon enzymic hydrolysis. The tetrasaccharide 7 (kcat/KM = 30000 M-ls 1) reacts 18-fold faster than the 4methylumbelliferyl trisaccharide 3c (kcat/KM = 1700 M'ls'l), equivalent to a difference in transition state activation energy (AAG~) of 1.9 kcal-mol 1. This difference in kcat~M is mainly due to kcat since both substrates have approximately the same KM values. If KM reflects the dissociation constant of the enzyme-inhibitor complex, this result suggests that the interaction of the glucopyranose unit of 7 with subsite +I is utilized in stabilizing the enzyme transition state with no additional stabilization of the enzyme-substrate complex.

OH

6

HHO ~

HO~a~,,,~.~ O OH

OH

3c

_OH

00~OxHO~Ox

_OH

_OH ~

:

OH

no hydrolysis

O.~. IVle

_OH o

_

Ho..L.,.,/~c~ HO4W,,,,'K,,-~O--~,~.'~,,--k..~-O~U-.~O

("OH

OH

OH

r OH

OH

(,.OH

~

OH

"IV "111 "11 "1 ' I Figure 6. Binding mode and kinetic parameters for substrates 3c, 6 and 7.

K M = 2.7 mM

kcat = 4.58 s1

KM= 2.6 mM

k at-7 s'

95 In terms of the enzyme mechanism, retaining glycosidases are believed to operate through a double displacement mechanism assisted by general acid-base catalysis [12]. Following the protonation of the glycosidic oxygen by the general acid, the mechanism involves some development of a positively charged oxocarbonium that is stabilized by the catalytic nucleophile either by electrostatic stabilization or by formation of a covalent glycosyl-enzyme intermediate. Since 7 and 3c only differ in the aglycon (leaving group) both leading to the same intermediate, the lower kcat value for 3c indicates that the rate determining step in the hydrolysis of the 4-methylumbellyferyl 13-glucan oligosaccharides is the initial glycosidation step up to the glycosyl-enzyme intermediate, prior to its hydrolytic break down by a water molecule.

3. ACKNOWLEDGEMENTS The authors thank Udo Heinemann (MCD, Berlin) for the resolution of the x-ray structure of the Bacillus licheniformis enzyme. This work was supported by Grant BIO94-0912-C02-02 from Comisi6n Interministerial de Ciencia y Tecnologfa (CICYT), Spain. C.M. acknowledges a fellowship from Generalitat de Catalunya.

4. REFERENCES

1 2 3 4 5 6 7 8 9 10 11 12

P.B. Hc~j, R. Condron, J.C. Traeger, J.C. McAuliffe and B.A. Stone, J. Biol. Chem., (1992) 267, 25059. F.W. Parrish, A.S. Perlin and E.T. Reese, Can. J. Chem., (1960) 38, 2094. B.A. Stone and A.E. Clarke, in Chemistry and Biology of (1---~3)-]3-Glucans, La Trobe University Press, Australia, 1992. A. Planas, M. Juncosa, J. Lloberas and E. Querol, FEBS Lett., (1992) 308, 141. M. Juncosa, J. Ports, T. Dot, E. Querol and A. Planas, J. Biol. Chem., (1994) 269, 14530. T: Keitel, O. Simons, R. Borriss and U. Heinemann, Proc. Natl. Acad. Sci. U.S.A., (1993) 90, 5287. T. Keitel, M. Meldgaard and U. Heinemann, Eur. J. Biochem., (1994) 222, 203. C. Malet, J. Jimdnez-Barbero, M. Bemab6, C. Brosa and A. Planas, (1993) Biochem. J., (1993) 296, 753. M.A. Anderson and B.A. Stone, FEBS Lett., (1975) 52, 202. C. Malet, J.L. Viladot, A. Ochoa, B. G~illego, C. Brosa and A. Planas, Carbohydr. Res., in press (1995). J.D. Allen, Methods Enzymol., (1980) 64, 248. M.L. Sinnott, (1990) Chem. Rev., (1990) 90, 1171.

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S.B. Petersen, B. Svensson, and S. Pedersen (Eds), CarbohydrateBioengineering 9 Elsevier Science B.V. All rights reserved.

97

Probing of glycosidase active sites through labeling, mutagenesis and kinetic studies Stephen G. Withers Protein Engineering Network of Centres of Excellence and Department of Chemistry, University of British Columbia, Vancouver, British Columbia, Canada, V6T 1Z1 Abstract Glycosidases which hydrolyses their substrates with net retention of anomeric configuration (retaining enzymes) do so via a double-displacement mechanism in which a covalent glycosylenzyme is formed and hydrolysed with acid/base catalytic assistance via oxocarbenium ion-like transition states. Strategies for identification of the nucleophilic residue and the acid/base catalytic residue have been devised. By use of mechanism-based inhibitors to trap the glycosylenzyme intermediate and affinity labels to derivatise the acid/base catalyst, in combination with novel mass spectrometric techniques, the identities of these residues have been determined in several glycosidases. In all cases identified to date these have turned out to be the carboxylic amino acids glutamate and aspartate. Armed with this knowledge, a retaining glycosidase has been converted into an inverter through replacement of the nucleophilic glutamate by alanine, along with the addition of alternative anionic nucleophiles such as azide and formate to the reaction medium. This provides the first example in which such a change of mechanism in a glycosidase has been effected.

1. INTRODUCTION The principal criterion by which glycosidases can be classified mechanistically is the stereochemical outcome of the bond cleavage which they carry out. One group of glycosidases hydrolyses the glycosidic linkage with net inversion of anomeric configuration, and these are known as inverting enzymes. Thus an inverting o~-glucosidase, for example, would hydrolyse an ot-glucoside substrate yielding ~3-glucose as the first-formed sugar product. Mutarotation would of course subsequently equilibrate this to the thermodynamic mixture of or- and ]3glucose. The second group of glycosidases hydrolyses the glycosidic linkage with net retention of anomeric configuration, and these are known as retaining enzymes. Different mechanisms are required for such different stereochemical outcomes, and indeed the fundamentals of these mechanisms were proposed over 40 years ago by Koshland [1]. These are shown in the diagram below in which the enzyme active sites are represented by boxes containing two key carboxylic acid residues. Clearly a large number of other residues are essential for specificity and catalysis, particularly those whose function is to bind to and stabilise the substrate as it

98 approaches its transition state conformation. However these have been omitted from this diagram in the interests of clarity.

RETAINING MECHANISM H

I

o

,fo

OH

HOiO

H

INVERTING MECHANISM

oLo H

OH

I

As is suggested by this diagram, there are considerable similarities between these two classes of enzymes in that both contain a pair of carboxylic acid residues suitably disposed on either side of the bond to be cleaved. However, as will become apparent, the roles of these two residues are different. In the case of the inverting enzymes, the mechanism involves a direct displacement in which the water attacks at the anomeric centre with general base catalytic assistance from one of the carboxylates. In a somewhat symmetrical manner, the other carboxylic acid functions as a general acid catalyst, protonating the leaving glycosidic oxygen in a process concerted with bond breaking. This results in hydrolysis with net inversion of anomeric configuration. The mechanism for the retaining enzymes, by contrast, involves a double displacement reaction in which a covalent glycosyl-enzyme intermediate is formed and hydrolysed. In the first step the deprotonated carboxylate functions as a nucleophile, attacking at the anomeric centre and displacing the glycosidic oxygen in a process which occurs with general acid catalytic assistance from the protonated carboxylic acid. This results in the formation of a covalent glycosyl-enzyme intermediate. In the second step water attacks at the anomeric centre of the glycosyl-enzyme species with general base-catalytic assistance from the deprotonated carboxylate, displacing the enzymatic active site nucleophile and releasing product sugar and free enzyme.

99 Both these steps, as well as the single step for the inverting enzyme, proceed via transition states with considerable oxocarbenium ion character. This means that, formally, bond cleavage precedes attack by the nucleophile, reaction proceeding via a dissociative (Snl) pathway. However, there is considerable evidence that there is still some degree of pre-association by the nucleophile, be it water or the active site carboxylate, presumably as one way of helping to stabilise the relatively unstable oxocarbenium ion [2]. The best evidence for the involvement of these oxocarbenium ion-like transition states has come from secondary deuterium kinetic isotope effect studies which provide insight into changes in hybridisation state at the anomeric centre between the ground state for the step in question and its subsequent transition state. Isotope effects between 1.05 and 1.25 have been measured for both steps (glycosylation and deglycosylation) for [~-glycosidases [3,4], and recently large isotope effects have been measured for ~-glycosidases [5]. Additional evidence has come from the potent inhibition afforded by inhibitors such as glyconolactones and amidinosugars which bear considerable structural resemblance to the proposed transition state [6]. Such inhibitors are, of course, recruiting the strong binding interactions which have evolved for stabilisation of the transition state conformation of the cleaving substrate. While the mechanism of retaining glycosidases is most conveniently described as proceeding through a covalent glycosyl-enzyme intermediate via oxocarbenium ion-like transition states, it is also quite possible that the oxocarbenium ion species flanking this covalent intermediate could be sufficiently stable to exist as short-lived species in their own right. This does not, however, significantly change the mechanistic description.

2. CARBOXYLIC ACIDS PLAYING A ROLE IN CATALYSIS 2.1. Identification of the active site carboxylic acids in retaining glycosidases Knowledge of the identities of the two key carboxylic acid-containing amino acids (aspartate or glutamate) in glycosidases is of considerable importance not only academically, but also for any structure/function or mutagenic studies as well as in cases where mutants are being made to provide enzymes of altered properties. A very effective, though typically rather labor-intensive, way of getting this information, as well as insight into the identifies of the other active site amino acids is through X-ray crystallographic determination of the 3-dimensional structure of the protein. Indeed, the structures of a number of glycosidases have now been determined crystallographically, and a compilation of the recently determined structures is provided in a recent review [7]. Frequently, however, this technique is not applicable, or available, and it is necessary to resort to other methods of identification. Further, even when a three-dimensional structure is available it is not always clear which amino acids are important unless the structure of an enzyme/substrate or enzyme/inhibitor complex has been determined. Even then it is also possible that the substrate or analogue may be bound in an unproductive mode, leading to a misassignment. An alternative approach involves the use of affinity labels, or far preferably, mechanismbased inactivators, to specifically derivatise the key amino acids in the active site. This requires the development of reagents which are capable of selectively reacting with and covalently "marking" the residues of interest. This manuscript reviews methods developed in the author's laboratory for the identification of these residues. Identification of active site nucleophiles in

100 glycosidases is carried out using a class of mechanism-based inactivator which traps the glycosyl-enzyme intermediate, in conjunction with electrospray tandem mass spectrometric methods for the localisation and sequencing of the labeled peptide within proteolytic digests. Identification of the acid/base residue can sometimes be achieved using affinity labels, or via detailed kinetic analysis of mutants modified at conserved glutamic and aspartic acids. 2.2. Identification of the active site nucleophile As noted above, the mechanism for retaining glycosidases involves the formation of a glycosyl-enzyme intermediate in which the sugar is covalently attached to the protein via the carboxylic side chain of a glutamic or aspartic acid. One way of identifying the nucleophilic amino acid residue would be to trap this intermediate by somehow decreasing its deglycosylafion rate. This requires differential manipulation of the rates of formation (glycosylation) and hydrolysis (deglycosylation) of the intermediate to slow the deglycosylafion step enormously from its typical values (t 1/2 around 1-10 ms) without ultimately slowing the glycosylafion step to as great an extent, since if this step were also slowed enormously, no intermediate would accumulate. This was achieved through the use of 2-deoxy-2-fluoro glycosides with good leaving groups such as dinitrophenolate or fluoride [8,9]. The presence of the C-2 fluorine substituent slows both the glycosylation and deglycosylation steps in two ways. One way relates to the fact that the hydroxyl substituent at C-2 plays a crucial role in transition state stabilisation in glycosidases by making key interactions (worth more than 8 kcal/mol), probably predominantly hydrogen bonding interactions, with the enzyme active site [10,11]. Replacement of the hydroxyl by fluorine, a substituent of limited hydrogen bonding potential, likely deletes, or at least decreases the value of, these hydrogen bonding interactions, destabilising both transition states significantly. The second way in which the fluorine substituent slows the two steps is through inductive destabilisation of the two electron-deficient transition states. Fluorine is much more electronegative than a hydroxyl, thus the positive charge developed at the transition state will be significantly destabilised by the presence of this substituent, slowing down both steps. The consequence of these two effects combined is a massive (up to 106 - 107 fold) reduction in rates of both steps [12,13]. Incorporation of a good leaving group such as 2,4-dinitrophenolate or fluoride speeds up the glycosylation step relative to the deglycosylation step with the effect that the intermediate is accumulated. Incubation of the enzyme with its corresponding 2-deoxy-2-fluoroglycoside then results in time-dependent inactivation, via the accumulation of a relatively stable 2-deoxy-2-fluoroglycosyl-enzyme intermediate.

H~

r

~1"~\ Ho4

-

o

HO/~O O~N

-o,r I

IH

.OH HO/~O o

,co I

o2N

o"

I

101 Supporting evidence for this mechanism of inactivation has been obtained in several ways. Firstly, stoicheometric reaction of inhibitor and enzyme has been demonstrated by electrospray mass spectrometry and by measurement of the magnitude of the "burst" of dinitrophenolate released [13]. Secondly, 19F-NMR studies of the inactivated enzyme have demonstrated the formation of a covalent ot-D-glycopyranosyl-enzyme intermediate [14]. Thirdly, and very importantly, the intermediate has been demonstrated to be catalytically competent. This was achieved by purifying inactivated enzyme of contaminating inactivator, then monitoring the reactivation of the enzyme consequent on turnover of this intermediate and release of the free enzyme [13,15]. Reactivation can be greatly accelerated by the inclusion of a suitable sugar acceptor into the reactivation mixture, such that turnover occurs via transglycosylation, a reaction typical of glycosidases. Identification of the amino acid residue labelled has been achieved in two distinct ways. The first of these required the synthesis of a radiolabelled version of the inactivator to generate a radiolabelled enzyme. Standard methods of proteolysis, HPLC separation of the resultant peptide mixture, and purification and ultimately sequencing of the radiolabeled peptide were then employed. This strategy was used successfully to identify the active site nucleophiles of four glycosidases, the ~-glucosidase from Agrobacterium faecalis, the exo-xylanase/glucanase from Cellulomonas fimi, the [3-galactosidase from Escherichia coli, and endo glucanase C from Clostridium thermocellum [15-18]. An alternative strategy, which obviates the need for synthesis of radiolabelled versions of the inactivator, involves the use of electrospray tandem mass spectrometry to identify the labeled peptide within a proteolytic digest, by monitoring for a collision-induced fragmentation reaction that is specific to the sugar-peptide linkage. This is illustrated below and in Figure 1 using the identification of the catalytic nucleophile in human [3-glucocerebrosidase by inactivation of the enzyme with 2-deoxy-2-fluoro-[3-glucosyl fluoride as an example [19]. Human 13-glucocerebrosidase is a lysosomal enzyme that cleaves the [3-glucosidic linkage of glucosylceramide. It is a membrane-associated glycoprotein (67 kDa, 497 amino acids) with both high mannose and complex oligosaccharides [20], whose crucial role in glycolipid catabolism has been established by disruption of the GCase gene in mice. Such mice die within hours of birth [21]. Inherited deficiencies of the human enzyme result in the variants of Gaucher disease, the most prevalent lysosomal storage disorder, affecting some 20,000 to 30,000 individuals worldwide. As such, this disease has been a prototype for the development of enzyme replacement therapies using specifically oligosaccharide-modified enzyme for targetting to the major sites of pathologic involvement, the macrophages of the liver, spleen and bone marrow [22-24]. Given the clinical, and commercial, importance of this enzyme, considerable interest in the identifies of active site residues is warranted, especially given the desire to understand the reasons for the inactivity of the enzyme in naturally occurring mutants. Treatment with 2-deoxy-2-fluoro-~-glucosyl fluoride (2FGlcF) resulted in time-dependent inactivation of the enzyme according to pseudo first order kinetics, yielding a value for the second order inactivation rate constant of ki/Ki = 22.7 min-lM 1. Inclusion of castanospermine (8.3 ~M), a known competitive inhibitor of GCase (Ki = 7 t.tM) in an inactivation mixture containing 10.4 mM 2FGlcF reduced kobs, the pseudo-first order inactivation rate constant, from 0.23 n ~ -1 to 0.15 min -1, showing that the inactivation was a consequence of reaction at the active site. Further the intermediate was shown to be catalytically competent by removal of excess inactivator from the labeled enzyme and then monitoring the return of activity. This was

102 a first order process, with a spontaneous reactivation rate constant, kreac of 5.3 x 10-4 min -1, corresponding to a half-life of tl/2 = 1300 minutes. The site of labeling was then identified by proteolysis of the labeled enzyme followed by HPLC separation, using an electrospray ionisation mass spectrometer (ESMS) to identify the labeled peptide by observing a specific, predicted fragmentation reaction, as follows. Peptic hydrolysis of 2FGlc-labeled GCase resulted in a mixture of peptides which was separated by reversed phase-HPLC using the ESMS as detector. When the spectrometer was scanned in the normal LC/MS mode, the total ion chromatogram (TIC) of the 2FGlc-labeled GCase digest displayed a large number of peaks, which arise from every peptide in the mixture (Fig. la). The 2-fluoroglucosylated peptide was then identified in a second run by using the tandem mass spectrometer set up in the neutral loss mode. In this technique the ions are subjected to

!

'| c i"1

IO

lo

~.1oo

J-

1~

14

111

18 INn.

10

~1oo

J:

I"''

12

14

10

18 mlrL

D

o

m~z

1000

11~00

Figure 1. ESMS experiments on glucocerebrosidase proteolytic digests.

limited fragmentation by collisions with an inert gas (Ar) in a collision cell located between the two quadrupole mass filters. Since the ester linkage between the sugar inhibitor and the peptide is quite labile under these conditions, relatively facile homolytic cleavage of this bond occurs with loss of a neutral sugar residue of known mass (165 Da) leaving the peptide moiety with its original charge. The two quadrupole analysers on either side of the collision cell can then be scanned in a linked manner in which the scanning of the two analyzers is offset by the mass of the anticipated "lost" neutral species. Under these conditions only ions differing in m/z by the mass of the lost sugar moiety (165 Da) can pass through both quadrupoles and be detected. As can be seen in Figure l b, when the spectrometer was scanned in this neutral loss tandem MS/MS mode searching for the mass loss m/z 165, corresponding to the loss of the 2FGlc label

103

100 ~

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-

50

m

FASEA+2r--Gk:

F~

25

FASE

11

3OO

4OO

" m/z

~/ 35OO

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7OO

Figure 2. Tandem MS/MS daughter ion spectrum of the 2FGlc-labeled peptide.

a true reflection of the activity of the enzyme, but results from contamination by wild type enzyme likely due to translational misincorporation, since equivalent levels of misincorporation have been observed previously [26,27]. Indeed, the activity observed is completely inactivatable by 2FGlcF at rates comparable to those of wild type enzyme, a result which is inconsistent with the expected enzymatic activity of a mutant in which the attachment site has been removed. This method has now been applied to a number of different glycosidases, including some whose catalytic nucleophile had been identified previously, namely Agrobacterium faecalis ~5glucosidase, Cellulomonas fimi exo-xylanase/glucanase and Clostridium thermocellum endo glucanase C [28], as well as the Bacillus subtilis xylanase [29]. The importance of these residues to catalysis has been confirmed in each case by kinetic analysis of mutants modified at those positions, and their active site location has been confirmed by X-ray crystallographic analysis of two of the enzymes, the Cellulomonas fimi exo-xylanase/glucanase [30] and the Bacillus subtilis xylanase [31 ]. 2.3. Identification of the a c i d / b a s e r e s i d u e by affinity labeling Techniques for the identification of this residue are not as well developed, or as reliable, as those described for identification of the catalytic nucleophile. One method which has been successful in several cases involves the use of an affinity label based upon a sugar with a reactive anomeric N-bromoacetyl functionality. The hope with this class of labels is that the reactive N-bromoacetyl moiety will react preferentially with the active site carboxylic acid in whose proximity it is bound, as shown in the Scheme below. I

.o~--~\

.

.........

.j

HO,,~

-

0

l

o, .o o

~c.O I

O~c..O t

104 N-Bromoacetyl 13-glycosylamines were first used as affinity labels with Escherichia coli [3galactosidase [32], the labeled amino acid being identified as Met501. More recently we have described their use as affinity labels for both the Agrobacterium faecalis 13-glucosidase and the Cellulomonas fimi exo-xylanase/glucanase [33]. In the former case, mass spectrometric analysis of the enzyme labeled with N-bromoacetyl ~-glucosylamine revealed that, even though simple pseudo-first order kinetics of inactivation had been determined, the inactivated enzyme was labeled with at least three equivalents of inactivator (Figure 3). The picture is further complicated by the presence of two native [~-glucosidase species of masses 51,205 and 51,066, corresponding to species with and without an N-terminal methionine residue. The presence of this N-terminal methionine residue, due to incomplete processing at the higher expression levels employed, has previously been shown to have no effect upon kinetic parameters. However in the case of the Cellulomonas fimi exo-xylanase/glucanase, inactivation by Nbromoacetyl ]3-cellobiosylamine resulted in the addition of only a single equivalent of inactivator per enzyme, as shown by the mass increase of (Am = 382) in Figure 4. The multiple labeling seen for the [3-glucosidase precluded simple identification of the site of labeling which led to inactivation, but the single labeling seen for the C. fimi exo-xylanase/glucanase provided an ideal system for study. Since it seemed unlikely that an appropriate neutral loss fragmentation would be observed in proteolytic digests of the labeled enzyme the identification of the site of labeling was attempted via comparative analysis of HPLC/ESIMS profiles of peptic digests of the labeled and unlabeled enzymes, using the mass information to "realign" the profiles and thereby correct for the irreproducibility of the HPLC chromatograms. A search was thus made for a peptide which was present in the labeled sample, but absent from the unlabeled sample, which was also greater in mass by exactly the mass of the label (Am = 382) than another peak which was present in the unlabeled sample, but (ideally) absent from the labeled. After searching through all the peaks only one peptide was found which met these criteria. Figure 5 shows the composite mass spectrum of all the peptides eluting from the HPLC between 19 and 24 minutes, the region in which the peptide of interest eluted. As can be seen, a peptide of m/z = 1028 is present in the digest of the labeled protein (Figure 5B), but is not found in the equivalent digest of the unlabeled enzyme (Figure 5A). Correspondingly, a peptide of rrgz = 646, smaller by the mass of the label (382), is found in the digest of the unlabeled enzyme, but only at lower intensity in the digest from the labeled. Ideally this peptide would not be found in the sample of the labeled protein digest, but is presumably present because of incomplete labeling or, more likely, because of partial degradation of the labeled peptide. A search of the amino acid sequence of Cex revealed only nine peptides of this mass which could be generated from this protein. The peptide was then further purified by reverse phase (C-18) HPLC, using the mass spectrometer as the detector, and sequenced by the Edman degradation in collaboration with Dr. Ruedi Aebersold, using the modified Edman reagent developed by his group and a mass spectrometric detector [34,35]. The great advantage of this approach, in which a mass spectrometer is used to identify the product modified phenylthiohydantoins, is that the mass of the modified amino acid phenylthiohydantoin is directly obtained. Using this approach the sequence DVVNEA was determined for this peptide, where the 5 th cycle resulted in the release of a phenylthiohydantoin derivative of m/z = 822. This is exactly the mass predicted for the phenylthiohydantoin of Nbromoacetylcellobiosylamine-modified glutamic acid.

105 1

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Figure 3. Reconstructed glucosidase

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'

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921

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892

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Figure 4: Reconstructed ESMS of C. fimi labeled with N-bromoacetylglucosylamine, exo-glycanase, a) free, b) labeled.

13-

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47,400

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936 9621183 806 922

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0.0

650

700

750

800

850

900

950

1000

1050

m/z

Figure 5. Mass spectra of proteolytic peptides of C. fimi exo-glycanase, A) free, B) labeled.

106 Alignment of the sequence of this peptide with the amino acid sequence of the enzyme indicates that the modified residue corresponds to Glu127. Gratifyingly this residue was recently identified as the most likely candidate for the acid-base residue on the basis of a detailed kinetic analysis of mutants modified at conserved glutamic and aspartic acid residues [36]. In addition, this residue has very recently been located in the active site of the enzyme through X-ray crystallographic determination of the 3-dimensional structure [30]. In addition to the above, the putative acid site acid catalytic residue in the I]-glucosidase from Manicot esculenta has recently been identifed using N-bromoacetyl 13-glucopyranosylamine as affinity label, locating the labeled peptide through use of radiolabels [37].

2.4. Identification of the acid/base residue by kinetic analysis of mutants The alternative approach to identification of the acid catalytic residue involves performance of a detailed kinetic analysis of mutants which have been created in which strictly conserved glutamic and aspartic acids are, individually, replaced by alanine residues. The approach is illustrated with the exo-glucanase/xylanase from Cfimi (Cex). Cex belongs to a family of more than 20 enzymes capable of hydrolysing xylan and cellulose. Hydrolysis occurs with net retention of anomeric configuration [38], thus it is a 'retaining' glycosidase, following a double displacement mechanism [39]. Sequence comparisons permit the identification of a number of conserved amino acid residues in this family, among which are 6 glutamic and aspartic acids, the probable candidates for the acid/base catalyst as noted earlier. As expected, it is found that the catalytic nucleophile Glu233 identified using the fluorosugar approach [16] is indeed one of these conserved residues, thus establishing one of the others as the probable acid/base catalyst. The strategy used thus involves mutation of these conserved Glu and Asp residues to alanine, then detailed investigation of the kinetic properties of the mutants so generated. Such an investigation requires the application of several mechanistic tests, as described below. As noted previously, the residue in question will function as an acid catalyst in the first step (glycosylation) and as a base catalyst in the second (deglycosylation). Therefore deletion of this residue might be expected to slow down both steps. However if a substrate is used which has a very good leaving group not requiring acid catalysis for its departure, then in that case the first step may not be significantly compromised for the mutant, but the second step necessarily will be since this step is common for all substrates of a fixed sugar type. Mutants modified at Glu 127 do indeed show this behaviour as can be seen in the table below. Rates of the first step, glycosylation, were assessed through kcat/Km measurements since in this system kcat~rn represents the first irreversible step, which is indeed glycosylation. Rates of deglycosylation were obtained from kcat values of substrates for which deglycosylation was rate-limiting. As can be seen, for an excellent substrate such as 2',4'-dinitrophenyl cellobioside (pKa of leaving 2,4-dinitrophenol = 4.0) the first step (kcat/Km) is not significantly slowed by mutation, whereas for a substrate requiring some acid catalysis such as 4'-nitrophenyl cellobioside (pKa of 4'-nitrophenol =7.0) there is a significant rate reduction, and for substrates with leaving groups of very high pKa such as 4-bromophenol (pKa = 10) the rate was much lower again. These results are therefore completely consistent with the role of Glu127 as the acid/base catalyst. A second consequence of removal of the acid/base catalyst is that a small cavity, possibly capable of accomodating an anion, will be generated at the ~-face of the substrate adjacent to

107 Table 1 Kinetic parameters for h~,drolysis of various substrates by Cex and E 127 mutants Enzyme Substrate kcat (min "1) Km (mM) kcat/Km (min-lmM-1) Native Cex DNPC 419 0.06 6983 PNPC 677 0.53 1278 PBrPC 255 2.0 128 E127A DNPC 2.4 0.0003 7742 PNPC 2.3 0.025 92 PBrPC 4.0 x 10-2 1.9 2.1 x 10-2

the anomeric centre. Such a site could permit the binding and attack of a nucleophilic anion at the anomeric centre. If a substrate with a good leaving group, such as DNPC, is employed, the second step (deglycosylation) will be rate-limiting, thus reaction of the intermediate with an anion more nucleophilic than water, and one not requiring general base catalysis, will result in an increase in the steady state rate. Such is indeed found to be the case, as is shown in Figure 6. Increasing concentrations of sodium azide as exogenous nucleophile result in progressive increases in kcat values. As can be seen, much greater overall rate increases are seen with DNPC (200-fold) than with PNPC (8-fold), rates reaching a limiting value at higher azide concentrations. This plateauing of rates is clearly due to a change in rate-determining step. As azide concentrations are increased, so the deglycosylation rate increases, and the steady state rate with it, up to the point at which the rate of the deglycosylation step becomes greater than that of the glycosylation step. Beyond this point no further rate increases are observed. The lower maximal rate observed with PNPC than with DNPC is therefore consistent with the smaller glycosylation rate constant for the substrate with the poorer leaving group. This mechanism therefore requires that the reaction product in the presence of azide be ]3cellobiosyl azide. This was clearly shown by chemical synthesis of ]3-cellobiosyl azide and comparison of its chromatographic behaviour and 1H-NMR spectrum with that of the reaction mixture. The two were shown to be identical. Interestingly, as can be seen in Figure 6, values of kcat/Km are esentially invariant with azide concentration. This is also completely consistent with the proposed mechanism since the parameter kcat/Km reflects the first irreversible step in catalysis, formation of the glycosyl-enzyme intermediate, and this step is not affected by nucleophilic competition with the water. These data therefore combine to strongly suggest that Glu127 is indeed the acid/base catalyst in Cex. This is, of course, the same residue that was identified using the Nbromoacetylcellobiosylamine affinity label, and as has been noted earlier, it has been found in the appropriate location in the active site by X-ray crystallographic analysis [30]. This concurrence of findings lends great confidence to this approach to identifying the acid/base residue. 3. MODIFYING THE MECHANISM As noted in the introductory section of this manuscript, both inverting and retaining glycosidases have a pair of carboxylic acids present in their active site, playing crucial roles,

108

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Figure 6. Kinetic parameters for hydrolysis of PNPC (panel A) and 2,4-DNPC (panel B) by Cex Glu127Ala in the presence of various concentrations of sodium azide.

albeit these roles are somewhat different in the two cases. In addition, both classes of enzyme catalyse hydrolysis via oxocarbenium ion-like transition states. There are therefore considerable similarities between the two classes of enzyme. However one difference which would be expected would be that the two carboxylic acids should be further apart in the inverting enzymes than in the retainers since, in addition to the bound glycoside, a water molecule has to be fitted in between the two groups. These expectations are indeed conf'm'ned upon inspection of crystal structures of enzymes of both mechanistic classes. Such an analysis [7,27] revealed that the two carboxylic acids are approximately 57k apart (oxygen to oxygen) in retaining enzymes, but approximately 9.5]~ apart in inverting enzymes. This opened up the possibility of converting an enzyme from one mechanistic class to the other by changing the separation of their carboxylic acids. The first attempt to do this on the Agrobacterium faecalis [3-glucosidase, for which no three-dimensional structural information is available, involved simply the mutation of the nucleophilic carboxylate, Glu358, to an aspartic acid, thereby increasing the separation, but only by about 1/~ [40]. This mutant was indeed active, with an activity some 2500-fold reduced over wild type, but product analysis revealed that it still followed the retaining mechanism. Clearly a greater separation was necessary if the mechanisms were to be changed, but in the absence of 3-dimensional structural information it was not clear how this would be

109 achieved. A second alternative involved the complete removal of this residue, replacing it by alanine so that it could not follow a retaining mechanism. This could work well if general base catalytic assistance for the attack of water were not too important. Such a mutant was indeed constructed, but as is shown in the Table below, it was essentially completely catalytically inactive, with an apparent kcat value some 107-fold lower than wild type enzyme. Clearly general base catalysis is very important as has indeed been shown for several other naturally inverting glycosidases (see for example ref. 41). However, addition to this mutant of small nucleophilic anions which do not need general base catalytic assistance resulted in enormous regains of activity of up to 105 fold, most of the way back up to wild type activity.

Table 2 Kinetic parameters for hydrolysis of 2,4-DNP-glucoside by Agrobacterium ~-glucosidase and its Glu358Ala mutant. Enzyme + activators kcat (s -1) Km (mM) Native glucosidase 89 0.031 E358A 7 x 10-6 0.1 E358A + 2M azide 1.1 3.8 E358A + 4M formate 3.0 1.1

Further, analysis of the reaction product in the presence of sodium azide revealed that ctglucosyl azide was indeed being formed, presumably through the inverting mechanism shown below.

I 70 H

o~C~oH

I o/JC~oH

OH H O H o ~ ~

o

OH

L_ N3 ~H3

OH I N3

CH3 I

This provides the first example of a change in mechanism being intentionally engineered into a glycosidase, and thereby serves both as a reassurance that the mechanisms and active site structures of these two classes of glycosidase (inverting and retaining) might not be so

110 different, and that it is indeed possible to modify these enzymes in a defined way so as to alter their catalytic activities.

4. A C K N O W L E D G E M E N T S

The author would like to thank the many co-workers and collaborators who made this work possible. Their names are listed in the references provided. I also thank the Natural Sciences and Engineering Research Council of Canada and the Protein Engineering Network of Centres of Excellence for financial support.

5. REFERENCES

9 10 11 12 13 14 15 16 17 18 19 20 21

22

D.E. Koshland, Biol. Rev. 28 (1953) 416. N.S. Banait and W.P Jencks, J. Amer. Chem. Soc., 113 (1991) 7958. M.L. Sinnott, Chem. Rev., 90 (1990) 1171. J.B. Kempton and S.G. Withers, Biochemistry 31 (1992) 9961. Y. Tanaka, W. Tao, J.S. Blanchard and E.J. Hehre, J. Biol. Chem., 269 (1994) 32306. G. Legler, Adv. Carb. Chem. Biochem., 48 (1990) 319. J. McCarter and S.G. Withers, Current Opinion in Structural Biology, 4 (1994) 885. S.G. Withers, I.P Street, P. Bird and D.H. Dolphin, J. Amer. Chem. Soc., 109 (1987) 7530. S.G. Withers, K. Rupitz and I.P. Street, J. Biol. Chem., 263 (1988) 7929. J. McCarter, M. Adam and S.G. Withers, Biochem. J., 286 (1992) 721. R. Wolfenden and W.M. Kati, Acc. Chem. Res., 24 (1991) 209. I.P. Street, K. Rupitz and S.G. Withers, Biochemistry, 28 (1989