approaches to labeling and identification of active site residues in

Protein Science (1995), 4:361-372. Cambridge University Press. Printed in the USA Copyright 0 1995 The Protein Society

REVIEW

Approaches to labeling and identification of active site residues in glycosidases

STEPHEN G . WITHERS’ AND RUED1 AEBERSOLD2 ’ Department of Chemistry, University of British Columbia, Vancouver, British Columbia V6T 1Z1, Canada ’Department of Molecular Biotechnology, University of Washington, Seattle, Washington 98195

(RECEIVED August 25, 1994; ACCEPTED December 27, 1994)

Abstract

Glycosidases play a key role in a number of biological processes and, as such, are of considerable clinical and bio- technological importance. Knowledge of the identities of catalytically important active site residues is essential for understanding the catalytic mechanism, for enzyme classification, and for targeted bioengineering of glycosidases with altered characteristics. Here we review and discuss traditional strategies and novel approaches based on tandem mass spectrometry for the identification of the key active site residues in glycosidases.

Keywords: active site residues; glycosidases; tandem mass spectrometry

Glycosidases are a diverse group of enzymes that catalyze the hydrolysis of glycosidic bonds. They therefore play a central role in a number of biological processes that are of significant interest for biochemistry, medicine, and biotechnology. For example, lysozyme is present in egg white and human tears and cleaves bacterial cell wall polysaccharides, thereby serving as a relatively broad range bactericidal agent. This enzyme, the first for which a detailed 3D structure was determined, thus has become the “prototypical” glycosidase and has served as a model enzyme for investigating structure, function, and catalytic mechanism of glycosidases.

The clinical relevance of glycosidases has become apparent, for example, through linkage of severe inherited disease phenotypes to defects in glycosidases responsible for lysosomal catab- olism. Since the introduction of the concept of lysosomal storage disorders (Hers, 1965) linking a fatal condition called Pompe disease to the absence of lysosomal a-glucosidase, defects in numerous other lysosomal glycosidases have been shown to be correlated with diverse disease phenotypes (Neufeld, 1991). One such enzyme, glucocerebrosidase (glucosylceramidase, acid 0- glucosidase), which, if defective, causes Gaucher disease, has been used successfully to treat patients with Type 1 Gaucher disease by enzyme replacement therapy (Barton et al., 1990).

Reprint requests to: Ruedi Aebersold, Department of Molecular Bio- technology, University of Washington, FJ-20, Seattle, Washington 98195; e-mail: [email protected].

Abbreviations: 3D, three-dimensional; ESI/MS, electrospray ioniza- tion mass spectrometry; RP-HPLC, reverse-phase high performance liquid chromatography; MS/MS, tandem mass spectrometry; 2FXb, 2-deoxy-2-fluoro-xylobiosyl; DNPZFXb, 2.4-dinitrophenyl 2-deoxy- 2-fluoro-xylobiosyl.

The commercial use of glycosidases in the biotechnology industry has also dramatically increased in recent years. Specific enzymes are increasingly used for food processing (invertase for production of “invert” sugar, cellulases for fruit juice processing, &galactosidase for lactose reduction), for “bio-stoning” of denim textiles (cellulases), for bio-bleaching in the pulp-and- paper industry (xylanases), and for biomass degradation with the potential to convert solid biomass into liquid fuels (Cough- Ian & Hazlewood, 1993).

Whereas numerous gene sequences coding for glycosidases have been determined (Henrissat, 1991; Henrissat & Bairoch, 1993), progress in understanding the biochemistry and structure of these enzymes has been much slower. Any application of these enzymes, particularly where that application could be enhanced by engineering of the enzyme’s structure and activity, will benefit from knowledge of the identities of the amino acid residues present at the active sites, and particularly of those directly involved in catalysis. In this manuscript, we briefly review previous approaches for the identification of catalytic residues in the active sites of glycosidases, focusing on recent develop- ments in the authors’ laboratories. We then interpret the respec- tive results within the models of glycosidase catalysis and the 3D structures of selected enzymes.

Classification of glycosidases

The vast array of glycosidases identified to date has been classified based on amino acid sequence similarities and on the catalytic mechanism. To date more than 480 complete glycosidase sequences have been determined and classified based on sequence similarities. The most extensive such work to date (Henrissat &

361

3 62 S.G. Withers and R. Aebersold

Bairoch, 1993) assigned 482 complete sequences into 45 families, plus 7 “orphan” sequences with no other family members. Fur- ther, a number of the larger families contain subfamilies of even higher internal sequence similarity. Sequence comparisons have revealed the modular structures of many of those enzymes that cleave polymeric substrates. In particular, many of the polymer- degrading glycosidases comprise, in addition to a catalytic domain, at least one domain responsible for binding of the enzyme to its (frequently insoluble) substrate. These binding domains may also play more active roles in physically disrupting the substrate structures but do not have catalytic activity. Such sepa- rate domains have been seen, for example, in cellulases and in glucoamylases (Svensson et al., 1989; Gilkes et al., 1991; Kilburn et al., 1993). In more general terms, sequence alignments can be extremely useful in suggesting amino acid residues that may play key roles in catalysis or in maintenance of structure. How- ever, additional information is required before any conclusions can be drawn concerning the roles of individual residues.

Mechanistic classification segregates glycosidases into two major categories. Those that hydrolyze the glycosidic bond with net inversion of configuration are termed inverting enzymes, and those that do so with net retention of anomeric configuration are termed retaining enzymes. These different stereochemical out- comes demand quite different catalytic mechanisms, and thus are likely to have different active site structures. Indeed it has been shown (Gebler et al., 1992b) that enzymes within in a sequence- related family all operate via the same catalytic mechanism.

Catalytic mechanism of retaining and inverting glycosidases

Likely mechanisms for these two enzyme classes were proposed by Koshland (1953) more than 40 years ago and have largely stood the test of time. Although the two mechanisms are dis- tinctly different, Figures 1 and 2 illustrate that they d o retain a number of features in common. Inverting glycosidases are believed to function by a single-step mechanism in which a water molecule effects a direct displacement of the glycosidic leaving group from the anomeric center as shown in Figure 1. This displacement mechanism is general acid/base catalyzed, with one active site amino acid acting as the general base, helping to de- protonate the nucleophilic water molecule, and the other amino

acid acting as a general acid, protonating the departing oxygen atom in a concerted fashion as the bond cleaves. The reaction proceeds via an oxocarbenium ion-like transition state. This model is supported by 3D structures that are available for several different inverting glycosidases such as the cellulases from Trichoderma reesei (Rouvinen et al., 1990) and Thermomono- spora fusca (Spezio et al., 1993), soybean @-amylase (Mikami et al., 1993), and Aspergillus awamori glucoamylase (Aleshin et al., 1994). In all cases studied, it is apparent that these acids and bases are the carboxylic side chains of aspartic or glutamic acids. Other studies, particularly those based upon kinetic isotope effects (Matsui et al., 1989), Bronsted relationships (van Doorslaer et al., 1984), or effects of substitution by halogens (Liu et al., 1991), have provided evidence that the transition state for these enzymes has considerable oxocarbenium ion character.

Retaining glycosidases are generally believed to function through a double displacement mechanism in which a glycosyl- enzyme intermediate is formed and hydrolyzed via oxocarbenium ion-like transition states as illustrated in Figure 2. Again the reaction is facilitated by acid/base catalysis, but in this case it is probable that the same group plays both roles. Three-dimensional structural information, which is available for several such enzymes, including the “original” glycosidase structure hen egg white lysozyme (Phillips, 1967), several a-amylases (Qian et al., 1993; Larson et al., 1994), cyclodextrin glucanotransferase (Klein et al., 1992), and Bacillussubtilisxylanase (Campbell et al., 1993; Wakarchuk et al., 1994), supports this model. In all cases, it is apparent that the two residues involved, the nucleophile and the acid/base catalyst, are again the carboxylic side chains of glutamic and aspartic acid residues. However, in the absence of information on the orientations of these residues relative to bound substrates or analogues, it is often difficult to assign these roles to specific amino acids.

Identification of active site amino acid residues by sequence alignment

With increasing numbers of complete glycosidase sequences available and classified into sequence homology groups, the identification of active site amino acid residues based on sequence alignment has become more useful. Sequences aligned within a homology group are analyzed for conserved acidic res-

Fig. 1. Catalytic mechanism for inverting glycosidases.

Identification of catalytic residues in glycosidases 363

I

HO L

o b * / F

1

OH

I T H- -A HO- 5H

HC

Fig. 2. Catalytic mechanism for retaining glycosidases.

idues that are predicted to be located in the catalytic sites in retaining as well as inverting glycosidases. Although this method can suggest catalytic residues, in itself it is not conclusive because in most cases a number of conserved Glu and Asp residues are observed, and not all of these will be located in the active site or involved in catalysis. Further insights into the roles in catalysis are usually obtained through site-directed mutagenesis at the candidate residues followed by detailed kinetic analysis of the mutants produced. Numerous examples of this approach exist (see Svensson & Sogaard, 1993), but all too frequently only a cursory evaluation of catalytic behavior of mutant enzymes is performed, thereby undoubtedly missing important conclusions. Further complications can arise in such analyses from the contamination of mutant proteins of intrinsically low activity with wild-type enzyme, either through lack of care in purification, through translational misincorporation, or genetic reversion (Schimmel, 1989). Considerable care is needed to assure that low activities found with mutant proteins are indeed true activities of the pure mutant, thus likely highly mechanistically informa- tive, and not the result of artefactual contamination.

Identification of active site amino acid residues by 3D structure analysis

If 3D structural information at atomic resolution is available, the identities of active site amino acid residues can often be determined by detailed examination of the active site region. Conclusive identification of the active site residues frequently requires determination of the structure of an enzyme/inhibitor or enzyme/substrate complex. In these structures, the residues that are important to catalysis, either directly as acidlbase, as nucleophilic catalysts, or less directly through binding of the substrate in its ground state or at the transition state, will be immediately apparent. However, 3D structures are still not available for the vast majority of glycosidases. Even for those whose structures are known, obtaining structural information on en- zymelligand complexes can be difficult. Further, even when the identities of the residues in close spatial proximity to the substrate are known, their specific roles in catalysis frequently can- not be predicted. Therefore, verification of roles in catalysis of specific residues postulated by 3D structure analysis requires ki-

364 S.G. Withers and R. Aebersold

B

Fig. 3. Structures of affinity labels.

netic analyses of carefully conceived mutants preferably in con- junction with studies using specific mechanism-based reagents.

Identification of active site amino acid residues using group-specific labels

Group-specific labels are designed to form stable, covalent bonds with specific functional groups in proteins. As such they generally d o not discriminate between functional groups located in the active site and those at any other location on the protein. They are therefore the least specific reagents for the identification of active site residues and their use has frequently led to mis- leading results, with almost every conceivable derivatizable amino acid having been identified at some stage as being “essential’’ to catalysis in some glycosidase. For this reason, this approach will not be discussed here beyond stating that probably the most successful and least unreliable class of group-specific reagents for glycosidases has been the carbodiimides. For example, Woodward’s reagent K and EAC have been used to identify potentially important carboxyl groups at the active sites of cellulases (Tomme & Claeyssens, 1989) and xylanases (Bray & Clarke, 1994), respectively. Although inherently inconclusive, results obtained with such reagents can at least be used to identify potentially important amino acid residues whose possible function can then be further probed through careful kinetic analysis of mutants modified at those positions.

Identification of active site amino acids by derivatization with affinity labels

This class of reagent generally contains a sugar moiety, which provides specificity for the active site, plus an inherently reactive functionality for forming stable conjugates with the enzyme. The specificity of this class of reagents, which is substantially increased over group-specific reagents, can be further enhanced by comparing the results from labeling experiments performed in the presence and absence of competitive, reversible inhibitors that protect the active site residues from derivatization by the affinity label. Several types of affinity label have been used in studies of glycosidases and structures of the glucosyl derivatives of two of these are shown in Figure 3. N-bromoacetyl glycosyla- mines (Fig. 3A) have been used to inactivate Escherichia coli &galactosidase (Naider et al., 1972), Aspergillus wentii 0-glucosidase (Legler, ]!BO), Agrobacterium faecalis &glucosidase, and

Cellulomonas fimi exo-glucanase/xylanase (Black et al., 1993). However, only in the case of the E. coli 0-galactosidase has the amino acid labeled been identified. The residue identified was methionine 502, which was later shown not to participate in catalysis. The isothiocyanate (Fig. 3B) (Shulman et al., 1976) has been used to inactivate the @-glucosidase from sweet almonds, but no labeled residue has been identified using this inactivator.

Considerably more effort has gone into the exploitation of glycosyl epoxide derivatives for the identification of active site residues. The epoxide derivatives are considered to function as shown in Figure 4, where protonation of the epoxide (either specifically by the acid catalyst or nonspecifically by some other residue) generates a reactive species prone to nucleophilic attack. Lysozyme was one of the earliest enzymes tested with this class of affinity reagent. The 2,3-epoxypropyl glycosides of N-acetyl- glucosamine, chitobiose, and chitotriose were synthesized and shown to function as affinity labels for hen egg white lysozyme, with the longer inhibitors binding and inactivating most effi- ciently. Using a I4C-labeled chitobiose derivative, it proved possible to purify and subsequently sequence the radiolabeled, proteolytically derived peptide (Eshdat et al., 1973). The amino acid identified in this work was aspartic acid 52, the residue previously postulated to act as the nucleophile, or charge-stabilizer, in the catalytic mechanism. This assignment was subsequently confirmed by X-ray crystallographic analysis of the inactivated enzyme (Moult et al., 1973). Interestingly, by reduction of the ester linkage, it proved possible to selectively convert aspartic acid 52 to a homoserine residue (Eshdat et al., 1974). The enzyme modified in this way remained catalytically inactive, thereby providing an early example of “mutation” of active site residues in proteins.

Epoxyalkyl glycosides have also been used with a number of other glycosidases, including sweet almond and A . wentii 0- glucosidases (Bause & Legler, 1974). The approach has been extended to other polysaccharidases, a number of different epoxyalkyl cellobiosides having been used successfully for investigating active site residues in cellulases. In the first such work, Legler used these reagents to inactivate cellulases from Asper- gillus niger, A . wentii, and Oxyporus sp. (Legler & Bause, 1973), the highest inactivation rates being found with the 4’,5’-epoxypentyl cellobioside. Subsequent work by other groups focused on cellulases and other 0-glucan hydrolases isolated from Schi- zophyllum commune (Clarke & Strating, 1989), B. subtilis, Hor- deum vulgare (Bordier Hoj et al., 1991), 7: reesei (Macarron et al., 1993), and Bacillus macerans (Keitel et al., 1993). The active site residue in the B. macerans enzyme was identified as Glu 105 by X-ray crystallographic analysis of the inactivated complex (Keitel et al., 1993). Although the residue labeled in some cases has been the active site nucleophile, such an assignment is by no means assured given the inherent flexibility of the alkyl chain and the remoteness of the reactive center from the bound sugar.

A more refined version of epoxide inactivators is found in the conduritol epoxides, which incorporate an endocyclic epoxide within a cyclitol ring, which itself mimics the sugar ring. A wide range of these epoxides has been synthesized to “match” the specificities of a range of glycosidases. The structure of one such epoxide (Fig. 5A), along with those of several other related compounds (Fig. 5B,C,D) is shown.

Inactivation is believed to follow the binding of the inactivator in a mode resembling that of the corresponding sugar sub-


Fig. 4. Inhibitory mechanism of glycosyl epoxide derivatives.

strate. Protonation of the epoxide and concomitant attack by an enzymic nucleophile lead to formation of a covalent derivative much as shown for the exocyclic epoxides, but likely in a more specific manner. A closely related class of compounds is that of the aziridines. Thus, conduritol aziridine (Fig. 5B) was synthesized and shown to be an effective inactivator of both a - and @-glucosidases, whereas the galacto-aziridine (Fig. 5C) rapidly inactivated green coffee bean a-galactosidase. This approach has been applied successfully to a wide range of glycosidases (see Legler, 1990, for an excellent compilation). These compounds have also been used in vivo to inactivate specific glycosidases in mice. An example is the inhibition of glucocerebrosidase in lysosomes by conduritol B epoxide providing a possible animal model for Gaucher disease (Kanfer et al., 1982).

In general it was found (Legler, 1990) that @-glycosidases are inactivated much more effectively by conduritol epoxides than are a-glycosidases, likely because trans opening of the epoxide by the @-glycosidase leads to the preferred trans-diaxial product. Ring opening with the a-glycosidases necessarily yields the less- favored trans-diequatorial product. Based upon this proposed mechanism of action, it was quite reasonably expected that the residue labeled in retaining glycosidases would be the active site nucleophile. Indeed, in at least one case (Bause & Legler, 1974), this was confirmed by "trapping" the catalytic intermediate formed during hydrolysis of p-nitrophenyl 2-deoxyglucoside by the A . wentii @-glucosidase and demonstrating that the amino acid residue involved was identical to that labeled with the conduritol epoxide (Roeser & Legler, 1981). However, the use of conduritol epoxides has led to the misassignment of active site residues in several known cases. In E. coli @-galactosidase and human lysosomal 0-glucocerebrosidase, subsequent kinetic analysis of mutants modified at the position determined using these reagents failed to display the expected ablation of catalytic activity and therefore cast doubt upon the proposed role for the

A B

identified residues. Indeed reinvestigation of these enzymes using a new class of inactivators, the 2-deoxy-2-fluor0 glycosides (see below) resulted in the identification of a different residue in each case, and mutants modified at this position indeed exhibited the expected kinetic behavior (Gebler et al., 1992a; Mia0 et al., 1994a; Yuan et al., 1994). The "incorrect" labeling observed with the conduritol epoxides is probably a consequence of imperfect mimicry of the parent sugar substrate, particularly given the absence of the C-6 hydroxymethyl substituent present in the hexopyranoside substrates. This presumably allows alternative binding modes in the enzyme active site, thus reaction with other amino acid residues. Interestingly, a naturally occurring epox- ycyclitol containing this hydroxymethyl group, cyclophellitol (Fig. 5D), was recently isolated from Phellinus sp. and shown in preliminary screens to exhibit glycosidase inhibitory activity (Atsumi et al., 1990). Detailed kinetic analysis with the A . faecalis @-glucosidase (Withers & Umezawa, 1991) revealed time- dependent inactivation, leading to an irreversibly inactivated enzyme. Further, this inactivator was much more specific with respect to the glycosidases it could inactivate than was the corresponding conduritol epoxide. Subsequent sequence analysis of the isolated, modified peptides (S. Mia0 & S.G. Withers, un- publ.) revealed that this inactivator does indeed label the same active site residue as that identified as the nucleophile by other methods (Withers et al., 1990). Cyclophellitol has also been used to inactivate glucocerebrosidases in vivo, thereby creating an animal model for Gaucher disease (Atsumi et al., 1992).

Identification of active site amino acid residues by active site labeling using mechanism-based inhibitors

Mechanism-based inhibitors, defined as relatively chemically in- ert ligands that require mechanism-based activation in order to subsequently react covalently with the enzyme, have been used

C D

Fig. 5. Structures of conduritol epoxide (A) and related compounds (B, C , D).

366 S . C. Withers and R . Aebersold

HO

NH2 *2

Fig. 6. Structure and activation of glycosylrnethyl triazenes.

quite successfully with glycosidases. Indeed, the conduritol epoxides just described could, arguably, be considered in this cat- egory. Another class of mechanism-based inhibitor that requires protonation after binding is that of the glycosylmethyl triazenes introduced by Sinnott (Sinnott & Smith, 1976; Marshall et al., 1981). As illustrated in Figure 6, protonation subsequent to binding yields a reactive species that decomposes rapidly to generate nitrogen, an arylamine, and, most importantly, a highly reactive glycosylmethyl carbenium ion. This reactive electro- phile, generated at the active site, then reacts rapidly with nearby nucleophiles, thereby blocking the active site and inactivating the enzyme. This approach has been applied to a number of enzymes (Sinnott & Smith, 1976; Marshall et al., 1981; Mega et al., 1990). By use of a radiolabeled version of this inactivator, two active site amino acid residues were labeled and subsequently identified in E. coli (lac Z and ebg 6-galactosidase). These residues, Met 501 and Glu 461, had previously been identified using other labels (Fowler et al., 1978; Fowler & Smith, 1983).

A second class of mechanism-based inhibitor that has been applied is one wherein the enzyme binds and cleaves the glyco-

sidic linkage of an inherently unreactive glycoside, then releases an aglycone that rearranges into a highly chemically reactive species. Examples are shown in Figure 7.

The first example is a difluoroalkyl glycoside (Fig. 7A), which undergoes enzyme-catalyzed cleavage, releasing a fluorohydrin. This rapidly eliminates hydrogen fluoride, yielding an acyl fluoride that can react with the enzyme active site (Halazy et al., 1989). The second example is a difluorotolyl glycoside (Fig. 7B) whose aglycone, upon enzymatic release, again liberates hydrogen fluoride, but this time yielding a reactive quinone methide (Halazy et al., 1990). Interestingly, a natural product that functions as a time-dependent inactivator of glycosidases, salicortin, has been isolated from plants of the family Salicaceae (Clausen et al., 1990). The proposed mechanism of action of this compound also involves the generation of a quinone methide.

This class of inhibitors is interesting and will undoubtedly have applications, but such compounds are of limited use for mechanistic studies because they release a reactive aglycone, which has no significant affinity for the enzyme active site. Thus, if it does not react immediately upon generation, it may react

B

H:*o OMe f O H

I

OH

I

CF2 H

0

Fig. 7. Structure and mode of activation of difluoroalkyl glycoside (A) and difluorotolyl glycoside (B).


quite nonspecifically. No studies in which labeled residues have been identified with this class of compounds have yet been reported.

Specific labeling of the active site nucleophile

Recently, activated 2-deoxy-2-fluor0 glycosides were developed as a new class of inactivator that functions via the formation and accumulation of a relatively stable glycosyl-enzyme intermediate (Withers et al., 1987, 1988). The structure and mode of action of such inhibitors are illustrated in Figure 8. The presence of the fluorine substituent at C-2 slows both the glycosylation and deglycosylation steps in two ways. The first is due to the fact that the hydroxyl substituent at C-2 plays a crucial role in transition-state stabilization in glycosidases by forming key interactions (worth more than 8 kcal/mol), probably predomi- nantly hydrogen bonding interactions, with the enzyme active site (McCarter et al., 1992). Removal, or diminution, of these interactions by replacement of the hydroxyl with fluorine, a substituent of limited hydrogen bonding potential, destabilizes both transition states significantly. Indeed this was the basis for the previous use by Legler of p-nitrophenyl2-deoxy-glucoside to ac- cumulate a glycosyl-enzyme intermediate on the A . wentii 0-glucosidase (Roeser & Legler, 1981). He subsequently identified the labeled amino acid after denaturation trapping. The second way in which the fluorine substituent slows the two steps is through inductive destabilization of the electron-deficient transition states. Fluorine is much more electronegative than a hydroxyl, thus the positive charge developed at the transition state will be significantly destabilized by the presence of this substituent, with consequent retarding effects upon the rates of both steps. The consequence of these two effects combined is a massive (up to 106-107-fold) reduction in rates of both steps (Street et al., 1989, 1992). Incorporation of a good leaving group (aglycones

as “reactive” as 2,4-dinitrophenolate or fluoride can be attached to sugar acetal centers and form reasonably stable species) 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 therefore results in time-dependent inactivation, via the accumulation of a relatively stable 2-deoxy-2-fluoroglycosyl-enzyme intermediate.

Supporting evidence for this mechanism has been obtained in several ways. Stoichiometric reaction of inhibitor and enzyme have been demonstrated by ESI/MS and by measurement of the magnitude of the “burst” of dinitrophenolate released (Street et al., 1992). I9F-NMR studies of the inactivated enzyme have demonstrated the formation of a covalent a-D-glycopyranosyl- enzyme intermediate (Withers & Street, 1988). The formation of a unique covalent linkage to a specific amino acid has been demonstrated by identification and sequencing of the peptide (Withers et al., 1990; Tu11 et al., 1991; Gebler et al., 1992a; Wang et al., 1993; Mia0 et al., 1994a, 1994b) (see below). Finally, and very importantly, the catalytic competence of this intermediate has been demonstrated by measuring turnover of the inactivated enzyme via hydrolysis of the intermediate, to yield free sugar and enzyme (Withers et al., 1990; Street et al., 1992). This 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, as shown in Figure 8B.

These inactivators have been used to identify the active site nucleophiles in a number of glycosidases, as shown in Table 1. The strategy employed for the identification of the residues in the first four cases was the relatively traditional one of synthesis and use of a radiolabeled inactivator to generate radiolabeled enzyme, then HPLC separation of proteolytically derived peptides, guided by the presence of radioisotope, and finally se-

A inactivation

f 0 0 ‘ch

B reactivation via transglycosylation

Fig. 8. Structure and reaction of activated 2-deoxy-2-fluor0 glycerides.

368

Table 1. Sequences of active site nucleophile peptides in glycosidases

Enzyme Sequence Reference

A. faecalis (3-glucosidase YITENGA Withers et al., 1990 C. fimi exo-glycanase VRITELD Tu11 et al., 1991 E. coli (3-galactosidase ILCEYAH Gebler et al., 1992a C. thermocellum endo-glucanase YCGEF Wang et al., 1993 Human glucocerebrosidase FASEA Mia0 et al., 1994a B. subfilis xylanase YGWTRSPLIEY Mia0 et al., 1994b

quencing of the purified peptide. This has been superseded by the new technology described below, which was used for the last two enzymes in the table.

Tandem mass spectrometric localization of active site nucleophiles

The uses of group-specific, affinity-based, and mechanism-based inhibitors for labeling active site residues have in common that the success of the experiment critically depends on a sensitive, fast, and unambiguous method for identifying the labeled residues. Use of affinity labels to identify residues has been limited, primarily by the complex syntheses frequently required for the introduction of a radioisotope into the inactivator. In addition, this approach is time consuming and technically demanding.

To circumvent the limitations inherent in the use of radiolabeled reagents, we have recently developed a new MS/MS- based technique for the identification of the residues covalently modified by mechanism-based inhibitors. The method is sche- matically outlined in Figure 9. It involves covalent labeling of the enzyme with a nonradioactive, mechanism-based inactivator, proteolytic digestion of the labeled, inhibited enzyme, identification and purification of the labeled peptides by RP-HPLC- ESI-MS/MS, and subsequent sequencing of the labeled peptide, either by the Edman degradation or by MS/MS. Three criteria, either by themselves or in combination, were used to select the labeled peptide in the complex peptide mixture. First, the labeled peptide is characterized by a different retention time on the RP-HPLC column compared to its unlabeled counterpart. This mobility shift is used to screen the LC/MS data for the disap- pearance of a specific peptide ion within a selected time window upon inhibition of the enzyme. Second, labeling of the peptide induces a characteristic mass shift by the addition of the 2-fluoro-glycosyl moiety. This is used to screen the LC/MS data set for the appearance of a new peptide ion species upon inhibition of the enzyme. Third, we have shown that the ester bond can be broken by collision-induced fragmentation in the collision cell of a tandem mass spectrometer, resulting in the loss of a neutral fluoro-glycosyl species. This neutral loss is characteristic for the glycosylated peptide and distinguishes glycosylated peptides within the peptide mixture from nonglycosylated peptides. In the triple quadrupole mass spectrometer the peptide un- dergoing the mass shift caused by the neutral loss can be detected if the first and the third quadrupole are scanned coordinately, but offset by the mass of the eliminated neutral species (neutral loss scan). This mass spectrometric method represents a rapid, sensitive, nonisotopic, and conclusive alternative to the standard

S.G. Withers and R . Aebersold

I I Glycosidase

Inactivation of enzyme with mechanism-based inhibitor

I Labeled-glycosidase I

1 Enzymatic digestion

r

Sugar I

Time ESI-MS/MS (neutral loss)

Identify candidate peptides within enzyme sequence

labeled peptide

Purify and sequence peptide

Time

Fig. 9. Schematic of tandem mass spectrometry-based techniques for identification of catalytic active site residues.

radioactive method. Only subnanomole amounts of inhibited protein are required and the scope of the method can be extended to any protein that can be specifically modified with a reagent that can fragment in a defined way in the mass spectrometer. The technique is illustrated with the following two examples in which active site nucleophiles in glycosidases are identified.

Identification of the active-site nucleophile of a xylanase

The B. subtilis xylanase can be inactivated by accumulation of its 2FXb-enzyme intermediate upon treatment with the mechanism- based inhibitor DNP2FXb. Peptic cleavage of 2FXb-labeled


10 15 20 min

-100- - ae B Peptide 1

$ 75 . Neutral Loss MS/MS: Xyl + 2FXb Digest Doubly Charged

5 0 '

,x 75 Neutral Loss MS/MS: Xyl. Digest

E O 10 15 20 min

825.5: (peptide + 2FXb +2H)'+

I

f 75 1 MS spectrum of peptide 1

Peptide mass = 1384

- 50

m h

Fig. 10. Determination of active site of nucleophile of a xylanase.

xylanase resulted in a mixture of peptides that was separated by RP-HPLC using the ESI/MS as detector. A flow-splitting de- vice was placed between the column and the electrospray ion- ization ion source so that part of the flow was directed into the ion source of the mass spectrometer and part of the flow was directed to a fraction collector (Hess et al., 1993). The ratio of flows was calibrated so that 90% of the sample was collected for further analysis and 10% of the sample was directed into the mass spectrometer. When the MS was scanned in the LC/MS mode, numerous peptides were detected in the peptic peptide mixture (Fig. 10A). The peptide bearing the 2FXb label was identified in a second run of the same sample using the MS in the neutral loss scanning mode. When the instrument was set to screen for a loss of m/z 133.5, corresponding to the loss of 2FXb (MW = 267) from the doubly charged active site peptide, a single peptide measured at m/z 826 was detected (Fig. 10B,D), whereas no peak was detected if a preparation of the noninhib- ited peptide was treated in an analogous manner (Fig. 1OC). Be- cause the doubly charged peptide was detected, the molecular weight of the peptide conjugate was 1,650 Da and the mass of the unlabeled peptic active site peptide was 1,384 (1,650 - 267 + 1 H). Sequencing of the selected peptide identified a region around Glu 78 as the actual active site peptide among several isobaric candidate peptides of 1,384 Da that could potentially be derived from digestion of the xylanase. This sequencing was performed both via the Edman degradation and via further MS/MS analysis, as shown in Figure 11. Fragments corresponding to the (apparent) loss of the indicated amino acid residues from the C-terminus are clearly seen. X-ray crystallographic analysis of the xylanase clearly revealed Gln 78 in a suitable position with respect to a bound substrate to function as the catalytic nucleophile (Wakarchuk et al., 1994).

b5: 664 b6: 751

b & k g A b9: 107

b10: 1204

Identification of the active site nucleophile of human glucocerebrosidase

Identification of the active site nucleophile of human glucocerebrosidase further illustrates the method and shows that, in

b8

1 J L loOa I b9

1385

bl0

mh

Fig. 11. MS/MS daughter ion spectrum of the 2F-xylobiosyl-xylanase active site peptide.

L 400

370 S.G. Withers and R . Aebersold

10 12 i 4 16 le min

" rb li 1'4 1'6 min

.- ,x Lo 7 5 1 L

- 2 50

- 5 25 a

0

._ !2 m

10 12 14 16 18 min

Fig. 12. Determination of active site nucleophile in human glucocerebrosidase.

some cases, in addition to the neutral loss scan, secondary criteria are required to conclusively determine the identity of the labeled peptide. Analogous to the situation with xylanase, peptic digestion of the 2F-glucose inhibited glucocerebrosidase resulted in a mixture of peptides (Fig. 12A). In this case, however, neutral loss scans under conditions suitable for detection of the 2F-glucosyl-labeled peptide revealed two major peaks (peptide 1, peptide 2, Fig. 12B) plus a number of minor signals, of which peptide 2 and the minor peaks also were present in the sample derived from the unlabeled peptide (Fig. 12C). The peptide of interest therefore corresponds to peptide 1 (m/z 688, Fig. 12D), the others arising from nonlabeled peptides that un- dergo an equivalent fragmentation. This most likely involves the elimination of a phenylalanine residue (165 Da, same mass as 2F-glucosyl moiety) from several different peptides. The active site peptide (peptide 1, Fig. 12B) was purified as described above and further characterized by peptide sequencing using MS/MS. Results from the sequencing experiment are shown in Figure 13. The authenticity of the residue identified (Glu 340) as the active site nucleophile was verified by kinetic analysis of a purified enzyme, which was altered at this residue by site-directed mutagen-

0

I FASEA+ZFGlc I I

FA 306 FAS

FASEA 523

I

300 400 500 600 700 d Z

Fig. 13. MS/MS daughter ion spectrum of the 2F-glucose-labeled glucocerebrosidase active site peptide.

esis (Miao et al., 1994a). The result obtained with this method therefore corrects the previously published identity of the catalytic residue of that enzyme (Dinur et al., 1986).

Summary and perspective

Over the last decade the number of available complete sequences of glycosidases has increased enormously. This abundance of primary structural information has allowed the classification of the enzymes into groups based on sequence similarities and mechanistic considerations. The synthesis and application of specific enzyme inhibitors, in particular mechanism-based inhibitors, have allowed the identification of the catalytic residues of these enzymes and increased our mechanistic understanding. Combination of such mechanism-based inactivation with ESI/MS-based methods for the identification and sequencing of the labeled peptide has provided a rapid and reliable way of obtaining this information. These results have now been corrob- orated by 3D structural information in many cases (Jacobsen et al., 1994; Wakarchuk et al., 1994; White et al., 1994). Equiv- alent approaches could undoubtedly be applied to many other enzyme systems with equally useful results.

Acknowledgments

We thank the many people who helped perform this work and whose names have appeared on the original papers. The excellent editorial as- sistance of Inge van Oostveen is gratefully acknowledged. This work was supported by the Natural Sciences and Engineering Research Council of Canada, the Protein Engineering Network of Centres of Excellence, and the Canadian Genetics Network of Centres of Excellence. R.A. was the recipient of a Medical Research Council (MRC) of Canada scholarship.

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