Structural and mechanistic insight into N-glycanprocessing by endo-α-mannosidaseAndrew J. Thompsona, Rohan J. Williamsb, Zalihe Hakkib, Dominic S. Alonzic, Tom Wennekesd, Tracey M. Glostera,Kriangsak Songsrirotea,e, Jane E. Thomas-Oatesa,e, Tanja M. Wrodniggf, Josef Spreitzf, Arnold E. Stützf,Terry D. Buttersc, Spencer J. Williamsb,1, and Gideon J. Daviesa,1

aDepartment of Chemistry, University of York, Heslington, York YO10 5DD, United Kingdom; bSchool of Chemistry and Bio21 Molecular Science andBiotechnology Institute, University of Melbourne, 30 Flemington Road, Parkville, Victoria 3010, Australia; cOxford Glycobiology Institute, Department ofBiochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, United Kingdom; dLaboratory of Organic Chemistry, Wageningen University,6703 HB, Wageningen, The Netherlands; eCentre of Excellence in Mass Spectrometry, University of York, Heslington, York YO10 5DD, United Kingdom;and fInstitute of Organic Chemistry, Graz University of Technology, Stremayrgasse 9, A-8010 Graz, Austria

Edited by Chi-Huey Wong, Academia Sinica, Taipei, Taiwan, and approved November 28, 2011 (received for review August 9, 2011)

N-linked glycans play key roles in protein folding, stability, andfunction. Biosynthetic modification of N-linked glycans, withinthe endoplasmic reticulum, features sequential trimming and read-ornment steps. One unusual enzyme, endo-α-mannosidase, cleavesmannoside linkages internally within an N-linked glycan chain,short circuiting the classical N-glycan biosynthetic pathway. Here,using two bacterial orthologs, we present the first structural andmechanistic dissection of endo-α-mannosidase. Structures solvedat resolutions 1.7–2.1 Å reveal a ðβ∕αÞ8 barrel fold in which thecatalytic center is present in a long substrate-binding groove,consistent with cleavage within the N-glycan chain. Enzymaticcleavage of authentic Glc1∕3Man9GlcNAc2 yields Glc1∕3-Man. Usingthe bespoke substrate α-Glc-1,3-α-Man fluoride, the enzyme wasshown to act with retention of anomeric configuration. Complexeswith the established endo-α-mannosidase inhibitor α-Glc-1,3-deox-ymannonojirimycin and a newly developed inhibitor, α-Glc-1,3-isofagomine, andwith the reducing-end product α-1,2-mannobiosestructurally define the −2 to þ2 subsites of the enzyme. Thesestructural and mechanistic data provide a foundation upon whichto develop new enzyme inhibitors targeting the hijacking ofN-glycan synthesis in viral disease and cancer.

3D structure ∣ enzyme inhibition ∣ enzyme mechanism ∣ glycobiology ∣glycosidase

N-linked glycans are present on the majority of eukaryoticproteins and direct their folding and influence their stability.

These polysaccharide decorations play important roles in pro-cesses such as protein folding, targeting, antigenicity, and lectininteractions, with defects leading to cellular dysfunction (1). Aber-rant N-glycan composition, through either incorrect or incompleteprocessing, is associated with various conditions, including Alzhei-mer’s disease, congenital disorders of glycosylation, viral infection,and metastatic cancer progression (2–4). Alteration of N-glycanbiosynthesis in cancerous cells and by human pathogenic virusesrenders the various proteins involved in their biosynthesis andmodification therapeutic targets. N-glycans are built-up and then“biosynthetically” degraded in an orchestrated process involvingsynthetic enzymes (glycosyltransferases) and catabolic enzymes(glycoside hydrolases) (5) (Fig. S1). Efforts to control diseasesof glycoprotein biosynthesis have largely focused on the use ofinhibitors of the “biosynthetic” trimming glycoside hydrolases in-volved in the early stages of N-glycan remodeling, such as gluco-sidases I and II. However, the failure of these inhibitors to providean effective treatment of these conditions likely occurs, in part,because of a unique enzyme, endo-α-mannosidase, which cleavesmannoside linkages internally, within the first branch of an N-gly-can chain (6–8).

N-glycans are covalently bound to the side-chain nitrogen ofasparagine (Asn) residues in the consensus Asn-Xxx-Ser/Thr (9).Within the endoplasmic reticulum (ER), presynthesized 14-mer

glycans are cotranslationally transferred en bloc by the multipro-tein complex oligosaccharyltransferase from the glycophospholi-pid precursor, Glc3Man9GlcNAc2-diphospho-dolichol, to Asnresidues within nascent polypeptide chains (Fig. S1A). The initialprocessing steps of this 14-sugar glycan commence while theunfolded glycoprotein remains attached to the ribosome, andinvolves the sequential removal of the terminal α-1,2-glucose byglucosidase I and removal of both the subsequent and final α-1,3-glucose moieties by glucosidase II, a procedure normally requiredfor the formation of mature N-glycans (1, 5, 10) (Fig. S1B). Undernormal conditions, the final α-1,3-glucose residue represents acheckpoint in protein folding and quality control because mono-glucosylated immature N-glycans with a terminal α-Glc-1,3-α-Man-1,2-α-Man chain are ligands for the molecular chaperonescalnexin (CNX) and calreticulin (CRT) (11). The fate of newlysynthesized glycoproteins is determined by a molecular inspec-tion process that monitors their folding state. Correctly foldedproteins exit the CNX/CRT cycle with removal of the final α-1,3-glucose from the Man9GlcNAc2 polysaccharide followed byα-mannosidase I cleavage of a mannose residue from the firstbranch and translocation to the Golgi apparatus. Misfoldedproteins undergo cycles of deglucosylation and reglucosylation,catalyzed by luminal UDP-glucose-dependent glycoprotein glyco-syltransferase. Terminally misfolded proteins are extracted fromthe folding cycle in a process termed ER-associated degradationand are retrotranslocated to the cytosol, where they are ubiqui-tinylated and proteasomally degraded.

Endo-α-mannosidase (classified into Carbohydrate-ActiveEnzymes Database family GH99; refs. 12 and 13; www.cazy.org)provides a glucosidase I and II independent pathway for thematuration of N-glycans (14). Endo-α-mannosidase hydrolyzesthe α-1,2-mannosidic bond between the glucose-substituted man-nose and the remainder of the N-glycan, and acts on the struc-tures Glc1–3Man9GlcNAc2 as well as structures that have beentrimmed by mannosidases in the 6′-pentamannosyl branch,releasing Glc1–3-1,3-α-Man oligosaccharides (15) (Fig. S1C).Despite growing insight into the biosynthetic function and subcel-

Author contributions: A.J.T., R.J.W., T.D.B., S.J.W., and G.J.D. designed research; A.J.T., R.J.W., Z.H., D.S.A., T.W., T.M.G., and K.S., performed research; R.J.W., Z.H., T.M.W., J.S., A.E.S.,and T.D.B. contributed new reagents/analytic tools; A.J.T., R.J.W., J.E.T.-O., T.D.B., S.J.W.,and G.J.D. analyzed data; and A.J.T., S.J.W., and G.J.D. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Freely available online through the PNAS open access option.

Data deposition: The atomic coordinates and structure factors have been deposited in theProtein Data Bank, www.pdb.org (PDB ID codes 4acy, 4acz, 4ad0, 4ad1, 4ad2, 4ad3, 4ad4,and 4ad5).1To whom correspondence may be addressed. E-mail: davies@ysbl.york.ac.uk or sjwill@unimelb.edu.au.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1111482109/-/DCSupplemental.

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lular localization of endo-α-mannosidase (14), no structures havebeen reported and nothing is known about its catalytic mechan-ism. Endo-α-mannosidase therefore represents an importantenzyme for further study, with the ultimate goal of developingapproaches to treat diseases involving aberrant N-glycosylation.We present a structural and kinetic analysis of two GH99 endo-α-mannosidase enzymes from the enteric bacteria Bacteroides the-taiotaomicron (BtGH99) and Bacteroides xylanisolvens (BxGH99),the first structures for any GH99 enzyme. We show that theseprotein orthologs are endo-α-mannosidases active on glucosylatedN-glycans and perform catalysis with a net retention of anomericconfiguration. Structures of complexes obtained with two aza/iminosugar inhibitors reveal intimate details of the catalytic residues,allowing the proposal of a unique blueprint for catalysis and pro-viding a structural rationale for inhibition in both a mechanisticcontext and ultimately for the development of therapeutic agents.

ResultsActivity and Kinetics of GH99 Endo-α-Mannosidase. B. thetaiotaomi-cron and B. xylanisolvens GH99 enzymes display 42% and 41% se-quence identity, respectively, toHomo sapiens endo-α-mannosidase(sequence alignments in Fig. S2). In order to ascertain whetherthe Bacteroides enzymes were appropriate models of the humanenzyme, activity on GlcMan9GlcNAc2 was studied by mass spectro-metry. Both BtGH99 and BxGH99 catalyze the removal of a dis-accharide from GlcMan9GlcNAc2, with peaks observed for bothreaction products, yet have no activity on the unglucosylated sub-strate Man9GlcNAc2, indicative of conversion to Man8GlcNAc2through release of a disaccharide, likely GlcMan (Fig. 1A). We nextstudied the BtGH99-catalyzed hydrolysis of a fluorescently labeledmammalian glycan, Glc3Man7GlcNAc2 by normal phase HPLC(see SI Methods). BtGH99 shows biologically relevant activityagainst this substrate with Km and kcat∕Km values of 83 μM and2.6 s−1 mM−1, respectively (Fig. S3). These data are in good agree-ment with the Km value of 55 μM determined for the rat liverenzyme studied by Lubas and Spiro (16). Thus, BtGH99 possessesspecific α-1,2-mannosidase activity when the −1 position manno-side is modified by one or more α-1,3-linked glucose residues. Noactivity was observed on a range of aryl α-mannosides. To furtherprobe the activity of this enzyme, the activated substrate, α-gluco-

pyranosyl-1,3-α-mannopyranosyl fluoride (Glc-ManF), was synthe-sized (see SI Methods). Hydrolysis of Glc-ManF releases fluoride,which is monitored using a fluoride ion-selective electrode.BtGH99shows high activity against Glc-ManF with Km and kcat∕Km valuesof 0.48 mM and 9.9 s−1 mM−1 (Fig. 1B). Although fluoride detec-tion is constrained to a relatively limited pH range, further obser-vation of this reaction across the pH range 5.5–7.5 revealed theenzyme to be optimally active at approximately pH 7.0, consistentwith another bacterial homolog from Shewanella amazonensis (17).That Glc-ManF acts as a substrate although various aryl manno-sides do not, coupled with activity on GlcMan9GlcNAc2 andGlc3Man9GlcNAc2, confirms BtGH99 as an endo-α-mannosidasewith a requirement for a minimal α-1,3-linked disaccharide sub-strate and hence with obligate binding for catalysis in a −2 subsite(for subsite nomenclature see ref. 18).

Further evidence for subsite specificity and the requirementfor a minimal occupancy of the −2 and −1 subsites was obtainedthrough binding of two BtGH99 inhibitors. α-Glucopyranosyl-1,3-deoxymannojirimycin (Glc-DMJ) (Fig. 1C) is in widespread useas a specific endo-α-mannosidase inhibitor and was developed bymodification of the known exo-α-mannosidase inhibitor deoxy-mannojirimycin with an endo-α-mannosidase-targeting 1,3-linkedα-glucosyl residue (19, 20). Inspired by the success of this ap-proach, a new inhibitor, α-glucosyl-1,3-isofagomine (Glc-IFG)(Fig. 1C), was synthesized (SI Methods) from the broad-spectrumazasugar inhibitor isofagomine, by introduction of a 1,3-linkedα-glucosyl residue to capitalize upon beneficial binding interac-tions in the −2 subsite. Isothermal titration calorimetry (ITC)revealed that BtGH99 binds both Glc-DMJ and Glc-IFG, withKd values of 24 μM and 625 nM, respectively (Fig. 2D). The in-hibition data support the requirement for a 1,3-linked disacchar-ide occupying the −1 and −2 subsites, which is further supportedby analysis of crystal structures of inhibitor-enzyme complexes(see below).

Three-Dimensional Structures of BtGH99 and BxGH99. The crystalstructure of a selenomethionine derivative of the B. thetaiotaomi-cron GH99 endo-α-mannosidase enzyme was solved at a resolu-tion of 1.70 Å (Table S1). Crystal structures of the native BtGH99and a second bacterial homolog BxGH99 were solved by mole-

Fig. 1. Activity, kinetics, and inhibition of B. thetaiotaomicron GH99 endo-α-mannosidase. (A) MALDI-TOF analysis of BtGH99 action on GlcMan9GlcNAc2 (m∕z2,600.0); (Upper) without enzyme and (Lower) with BtGH99 to yield a Hex8GlcNAc2 product consistent with removal of Glc α-1,3-Man (the enzyme has noactivity on Man9GlcNAc2). (B) Michaelis–Menten kinetics of BtGH99 on the bespoke substrate Glc-Man fluoride (shown in inset). (C) The endo-α-mannosidaseinhibitors Glc-DMJ (synthesis in ref. 19) and Glc-IFG (synthesis in this work). (D) Isothermal titration calorimetry of Glc-IFG binding to BtGH99.

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cular replacement at resolutions of 2.00 and 1.90 Å, respectively.Together, the different crystal forms and asymmetric unit con-tents yield 10 crystallographically independent “views” of thestructure. The three-dimensional structures of these bacterialendo-α-mannosidase enzymes reveal a single-domain proteinadopting a ðβ∕αÞ8 barrel fold with a central open cleft formedfrom extended loop regions linking the secondary structuralmotifs comprising the catalytic active site, Fig. 2A. These loopregions adopt different conformations in the various structuresdetermined, and given the extended and branching nature ofthe N-glycan substrate(s), it appears that these large flexible loopsare capable of forming stabilizing interactions with the substrate,positioning the scissile α-1,2-mannosidic bond so as to promotecatalysis within the active site. Perhaps unsurprisingly given theunique enzymatic activity assigned to GH99, this loop-adornedbarrel motif appears unique, with both bacterial GH99 orthologsexhibiting no significant structural and sequence similarity toglycoside hydrolases from other families. Analysis against knownstructural motifs using the Dali server (21) reveals weak second-ary structure matches, the closest being endo-β-1,4-mannanasefrom Trichoderma reesei (Protein Data Bank ID code 1QNR,Z ¼ 15.9, rmsd 3.1 Å across 233 Cα positions, sequenceidentity ¼ 8%).

The GH99 active center, located at the terminus of a solvent-accessible channel near the center of the barrel fold, possesses acluster of carboxylate side chains likely to play a role in manno-sidic bond hydrolysis. In order to help assign potential catalyticfunction to these residues, a time-course of BtGH99-catalyzedhydrolysis of Glc-ManF was studied by 1H NMR spectroscopy.Enzymatic hydrolysis of the α-linked C–F bond resulted in therapid appearance of a product, shown by 2D NOESY analysisto be α-Glc-1,3-α-mannose, thus indicating that GH99-catalyzedsubstrate hydrolysis occurs with a net retention of anomeric con-figuration (Fig. S4). Over several hours, the initial product wasobserved to undergo mutarotation, isomerizing to an equilibriummixture of both α- and β-anomers at the reducing-end mannose.

Classical retaining glycoside hydrolases operate through a two-step, double displacement reaction in which a covalent intermedi-ate is formed and then hydrolyzed, via oxocarbenium-ion-liketransition states (glycosidase mechanisms recently reviewed inref. 22). In most cases, this displacement reaction is achievedby an enzymatic nucleophile such as a carboxylate (aspartate orglutamate) or, as for certain sialidases, a tyrosine residue. In some2-acetamidoglycosidases (including families GH18, 20, 25, 56, 84,and 85), the nucleophile is the 2-acetamido group of the sub-strate, which acts in a “neighboring group” participation reaction.For GH99, we initially assumed that the catalytic apparatus of theenzyme comprised a catalytic nucleophile acting to form thecovalent glycosyl-enzyme intermediate and a catalytic acid/base,first functioning as a general acid to assist leaving group depar-ture and then as a general base to activate water for nucleophilic

attack at the anomeric carbon of the intermediate. Kinetic ana-lysis of three BtGH99 active-center variants, E154A, E329A, andE332A (E156, E333, and E336 in BxGH99), reveals substantialdecreases in catalytic activity (Table S2). Both the E154A andE329A variants show near zero activity. E332A meanwhile showsreduced activity with observed rate constants indicating anapproximate 50-fold decrease in catalytic activity with respect towild type using the activated Glc-ManF substrate. Against thenatural substrate, Glc3Man7GlcNAc2, the same mutant showszero activity compared to wild type, under matching experimentalconditions. In order to gain insight into the functions of the dif-ferent components of the catalytic apparatus, 3D structures weredetermined with a bacterial endo-α-mannosidase in complexwith inhibitors and the truncated reducing-end product, α-1,2-mannobiose.

Active Site Structure and Catalysis. The apo-BtGH99 crystal formpresented a major obstacle to the observation of enzyme-ligandcomplexes, owing to a large loop region (residues 64–79) project-ing into the active-center of another molecule of BtGH99 formingan artefactual crystallographic dimer. Despite considerable effort,attempts to obtain a form of BtGH99 amenable to complex forma-tion proved unsuccessful. A second bacterial ortholog, B. xylanisol-vens endo-α-mannosidase, was therefore selected for structuralcharacterization and proved amenable to complex formation,allowing structural determination of binary complexes withGlc-DMJ and Glc-IFG, and ternary complexes with each inhibitorand the reducing-end product α-1,2-mannobiose (Fig. 3).

Imino and aza sugars have proved instrumental reagents inthe glycosidase field (23, 24) both for structural and mechanisticdissection and as templates for therapeutic agents. Deoxynojiri-mycin-type iminosugars possess a basic nitrogen in place of theendocyclic ring oxygen and are mimics of an oxocarbenium-ionbearing positive charge on the ring oxygen position. Isofagomine-type azasugars possess a basic nitrogen at the pseudoanomericposition, and can be considered mimics of a glycosyl cation withcharge located on the anomeric carbon. These compounds arelikely to prove effective inhibitors and high-affinity ligands ofthe various GH99 endo-α-mannosidase orthologs. The complexesof BxGH99 with Glc-DMJ (25) and Glc-IFG unveil the ligand–protein interactions within the −1 and −2 subsites (18) (Fig. 3).

Fig. 2. Three-dimensional structure and conservation in GH99 endo-α-mannosidases. BxGH99 in complex with Glc-IFG and α-1,2-mannobiose as aribbon (A) and (B) a surface representation colored by sequence conservation(40) using the partial GH99 alignment as shown in Fig. S2.

Fig. 3. Electron density and ligand binding to GH99 endo-α-mannosidase.A–C represent binding of (A) Glc-DMJ, (B) Glc-IFG, (C) Glc-DMJ/α-1,2-mannobiose (in divergent stereo). Figures shown are REFMAC maximum-likelihood/σA weighted 2Fo − Fc syntheses contoured between 0.26 and0.32 electrons perÅ3.

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Both inhibitors bind to the enzyme active site with the −1 ringlocated deep within a substrate-binding pocket in an undistorted4C1 conformation. The −2 glucosyl residue projects from thecatalytic cavity and appears solvent accessible. These subsites liein the center of a substrate-binding cleft with a length of approxi-mately 40 Å. The solvent-exposed nature of the −2 subsite gluco-side allows the accommodation of elongated substrates withadditional glucose residues at the 3 position. Aromatic platformsare common binding elements of carbohydrate active enzymes(26) and in these distal negative subsites are provided by a pairof highly conserved tryptophans at positions 48 and 126. Se-quence alignments reveal a high degree of sequence conservationacross various species in the region comprising the substrate-bind-ing cleft (Fig. 2B). Indeed, all the interactions in the −2 toþ2 sub-sites appear to be essentially invariant (Figs. S2 (alignment) andS5 (interactions)). That the glucoside moiety interacts with resi-dues in the −2 subsite, and also with the putative catalytic appa-ratus, provides a structural rationale for the catalytic requirementof a sugar occupying the −2 subsite of endo-α-mannosidase.

Aza and imino sugar inhibitors have been used to identify thecatalytic apparatus of glycosidases by virtue of the interactionsof potential nucleophiles with the protonated nitrogen atoms(25, 26). BxGH99 structures in complex with Glc-DMJ andGlc-IFG reveal a water molecule, coordinated by Glu336 (332 inBtGH99) and poised below the pseudoanomeric nitrogen ofGlc-IFG and the C1 of the DMJ moiety of Glc-DMJ, in the idealposition for nucleophilic attack to complete the second hydrolysisstep of a classical double displacement retention mechanism.Glu336 is situated on the “α-face” of the sugar occupying the −1subsite, in a position where it might interact with the glycosidicoxygen, thus fulfilling a role strongly indicative of the catalyticacid/base in the enzyme mechanism. Consistent with such afunction, the BtGH99 variant E332A retains approximately 2% ac-tivity, compared to WT, on the activated fluoride substrate but hasno detectable activity on the natural GlcMan9GlcNAc2 glycan (de-termined by mass spectrometry) or against Glc3Man7GlcNAc2(determined by HPLC).

Isofagomine-type azasugar inhibitors are useful crystallo-graphic probes that typically allow direct observation of the en-zymatic nucleophile of retaining glycoside hydrolases. Uponprotonation, the pseudoanomeric nitrogen forms a salt bridgewith the negatively charged nucleophile, acting as an “ionic trap”for the nucleophile, which during the reaction coordinate mustapproach within bonding distance of the anomeric carbon (23).All X-ray structures of retaining glycoside hydrolases with isofa-gomine-type compounds have, without exception, revealed thecatalytic nucleophile to be located within 3 Å of the azasugarnitrogen, whereas for inverting enzymes, the nucleophilic wateris found less than 3 Å away from nitrogen (Table S3). Of parti-cular interest is the absence, in the GH99 orthologs, of any en-zymatic carboxylate at a position suitable to act as the enzymaticnucleophile in a conventional double displacement reaction.Indeed, without invoking a conformational change, or reposition-ing of the −1 sugar residue, the BxGH99 complex structures showno protein residue in a position to act as a potential nucleophile;the most closely located candidates for a classical double displa-cement mechanism (shown in Fig. 4A) include the OE1 atom ofGlu333 (approximately 3.5 Å distant) or the OH of Tyr46 orTyr252 (4.0 Å distant). However, none of these residues appearclose enough to interact with the inhibitor nitrogens at either ringposition in the complexes (Movie S1). Furthermore, the observa-tion of identical (and static) arrangements of the active site re-sidues in three crystal forms of two enzymes with a total of 10different packing arrangements provides no evidence of intrinsicconformational flexibility within the active site. Provocatively, it iswell known that the alkaline solvolysis of α-mannosyl fluorideproceeds through a mechanism involving neighboring groupparticipation in which the 2-OH attacks the anomeric positionresulting in the formation of the intermediate 1,2-anhydro-β-mannopyranose. This intermediate is opened by a nucleophile toafford an α-configured product possessing a retained anomericconfiguration (27). Although no evidence currently supports theexistence of a 1,2-anhydro sugar as an intermediate for glycosi-dase-catalyzed hydrolysis, it is possible that such a mechanismmay occur with endo-α-mannosidase (Fig. 4B). Accordingly, the

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Fig. 4. Putative catalytic mechanisms for GH99 endo-α-mannosidase. (A) Classical two-step double displacement mechanism proceeding via a glycosyl-enzymeintermediate; in the case of the structures reported here, this would require conformational change. (B) Neighboring group participation via a 1,2-anhydrosugar intermediate.

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kinetically essential and conserved active-center residue Glu333,poised 2.6 Å away from the O2 group of the substrate, woulddeprotonate the 2-OH, promoting its nucleophilic attack on C1.The anti-arrangement of O2 and the leaving group seen in thecomplex with Glc-DMJ is that stereoelectronically required forepoxide formation, which requires a linear transition state ofthe attacking nucleophile, the reactive carbon center, and theleaving group. Neighboring group participation by the 2-acetami-do group of N-acetyl-β-glycosides is now well established infamilies GH18, 20, 25, 56, 84, and 85 (28). Together the precedentfor neighboring group participation in these glycoside hydrolasefamilies, the lack of an apparent enzymatic nucleophile in theGH99 structures, and the established neighboring group partici-pation mechanism for nonenzymatic hydrolysis of α-mannosylfluoride, provides an enticing precedent to suggest this unortho-dox enzymatic mechanism.

In order to dissect the leaving-group (“positive” numbers;ref. 18) subsites of the GH99 endo-α-mannosidase, additional crys-tallization experiments were conducted with either Glc-DMJ orGlc-IFG together with α-1,2-mannobiose. Such conditions affordcomplexes with the inhibitors in the −1 and −2 subsites, and withthe mannobiose disaccharide in the þ1 and þ2 subsites. As withthe −2moiety, theþ2mannose residue appears solvent accessible,projecting outside the active site cavity at a distance of approxi-mately 7 Å from α-1,3-glucose, consistent with an ability to accom-modate the remaining reducing end residues of a complexN-glycan. Phenylalanines at positions 253 and 258 provide anaromatic platform positioning the þ1 mannoside, with Tyr289and Asn298 making hydrogen-bonding interactions with the O3andO4 positions of the same sugar. Likewise within theþ2 subsite,hydrogen bonding occurs between O3 and O4 and Tyr195, Asp196,and Tyr198. Despite the sugar residue occupying the þ1 subsiteappearing to be slightly displaced from its likely true position dur-ing hydrolytic cleavage of the authentic substrate (O2 of the þ1sugar lies approximately 4 Å from the atom equivalent to theanomeric C1 of the residue in the −1 subsite in both inhibitor com-plexes), both structures show the carboxylate group of the likelycatalytic acid/base, Glu336, to be in close proximity to the positionof the scissile α-1,2-mannosidic bond.

DiscussionEndo-α-mannosidase is a resident of the cis/medial compartmentof the Golgi apparatus and pre-Golgi intermediates, and is a mem-brane-associated protein that is difficult to recombinantly expressin soluble form (29). The bacterial orthologs from B. thetaiotaomi-cron and B. xylanisolvens are soluble proteins and may have beenacquired by horizontal gene transfer because these organisms arecommon and beneficial components of the human gut (30). It hasbeen suggested that, under normal conditions, endo-α-mannosi-dase acts to deglucosylate folded Glc1Man7–9 glycoproteins thatmay reach the Golgi apparatus through being poor substratesfor ER α-glucosidase II (7). The biological role of the bacterialenzymes is unclear, but may include, as is the case for the exo-α-mannosidases of family GH92 (31), mannose foraging for basicmetabolic inputs. Just as incomplete deglucosylation in the mam-malian cell prevents the action of exo-α-mannosidases, a similarproblem with the bacterial exo-α-mannosidases might have ledto the beneficial acquisition of the enzyme by the bacteria. It isnotable that whereas the genomes of B. thetaiotaomicron and B.xylanisolvens possess many copies of the N-glycan active exo-α-mannosidases (24 and 17 GH92 enzymes, respectively; 3 GH125enzymes each) each genome contains only a single copy of a GH99enzyme, which supports its role in relieving a metabolic bottleneckin the degradation of glucosylated N-glycans. Alternatively, oradditionally, the bacterial endo-α-mannosidase may act on yeastmannans, which contain the epimeric α-mannosyl-1,3-mannosesubstructure. In this regard, it is interesting that studies of mam-malian endo-α-mannosidase have indeed shown that α-mannosyl-

1,3-deoxymannonojirimycin, the epimer of Glc-DMJ, is an effec-tive inhibitor (20).

Mammalian endo-α-mannosidase has a preference for mono-glucosylated N-glycans and its activity increases as the numberof mannose residues on the 6′-mannose branch decreases (16).The structures of the bacterial orthologs suggest that the majorityof substrate specificity is for the −1 and −2 subsites with addi-tional sugars off the 3-position of the −2 glucose residue likelyto project into solvent and not to be involved in significant en-zyme interactions. α-Mannosidases must overcome certain sub-strate-specific challenges to achieve effective catalysis becausethe enhanced anomeric effect in mannose deters substrate distor-tion, and thus an attacking nucleophile suffers destabilizing 1,2-diaxial interactions with the axial 2-OH group. In retaining GH38(32), and inverting GH47 (33) GH92 (31) α-mannosidases, adivalent metal ion (Zn2þ for GH38, Ca2þ for GH47 and GH92)is found to coordinate the 2- and 3-hydroxyls, possibly assisting incatalysis through substrate distortion and in delivery of the nu-cleophile (31). By contrast, in the inverting GH125 enzymesno metal ion is observed in any structures and no substrate dis-tortion was seen in a pseudo-Michaelis complex with an S-linkeddisaccharide spanning the active site (34). It is unclear how thisfamily overcomes the destabilizing diaxial interaction of the at-tacking water nucleophile, although it is possible that a favorablehydrogen bond between the 2-OH of the sugar and the nucleo-philic water may assist in nucleophilic attack in a similar way tothe divalent metals in the GH38, GH47, and GH92 enzymes. Anotable feature of the GH99 active site observed in both bacterialendo-α-mannosidase orthologs is the absence of a metal ion orindeed of a site for possible metal ion coordination. The en-zyme-inhibitor complexes with Glc-DMJ and Glc-IFG deter-mined for BxGH99 reveal no obvious substrate distortion in the−1 subsite and, despite the presence of an ionic trap in Glc-IFG,no enzymatic nucleophile is apparent. However, a widely con-served residue, Glu333, is found in close proximity to O2, andits mutagenesis results in a near-complete decrease in catalyticactivity. Taken together, these observations suggest that thehydrolysis reaction catalyzed by the GH99 endo-α-mannosidasefamily proceeds via both a metal-independent and possibly auniquely unorthodox mechanism.

Inhibitors that intervene in early steps of N-glycan biosynthesishave been largely ineffective in suppressing N-linked glycanprocessing. Cells in which glucosidase II has been inhibited orsubjected to genetic knockout retain 40–80% normal N-glycanprocessing function through the intervention of endo-α-manno-sidase (8, 35, 36). During a castanospermine (CST)-imposedα-glucosidase I and II blockade, HepG2 cells produced N-linkedglycoproteins with the normal glycan structure and resultedin concomitant release of free Glc1–3Man oligosaccharides (6).Evidence for the basal action of endo-α-mannosidase in the ab-sence of glucosidase inhibition or knockout was demonstratedthrough the use of the mannosidase I inhibitor, DMJ, which pre-vents further processing of the deglucosylated N-linked oligosac-charides. Man6–8GlcNAc, but not Man9GlcNAc, structures wereidentified, with the Man8GlcNAc structure being the character-istic isomer generated by endo-α-mannosidase cleavage (6).Coadministration of the α-glucosidase I and II inhibitor CSTandthe selective endo-α-mannosidase inhibitor Glc-DMJ resulted ininhibition of biosynthesis of Glc3Man and N-linked glycoproteins(25). On the other hand, Golgi α-mannosidase II is seen as apromising target for therapeutic intervention, as this enzyme actsafter endo-α-mannosidase rejoins the normal pathway and so actson all N-linked glycans undergoing processing (25). Swainsonine,a selective inhibitor of Golgi α-mannosidase II, blocks the abnor-mal formation of complex oligosaccharides, leading to reducedmetastasis and tumor growth (37). However, swainsonine alsoinhibits the closely related lysosomal α-mannosidase, limiting itsclinical use (38). Combination therapies of glucosidase I/II inhi-

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bitors (e.g., 6-O-butanoyl-CST) and endo-α-mannosidase inhibi-tors may provide a viable alternative approach. The developmentof Glc-IFG as a more potent inhibitor of the bacterial endo-α-mannosidase serves as a valuable precedent for glucosylationof inhibitors of exo-α-mannosidases to search for more active in-hibitors of endo-α-mannosidase, which will be assisted by thestructural and mechanistic blueprint provided by this work.

MethodsGene Cloning, Mutagenesis and Protein Production. The gene encodingBtGH99 was amplified from genomic DNA by PCR and ligated into a modifiedpET28a vector encoding an N-terminal His6-tag (pETLIC-BtGH99). Site-direc-tedmutagenesis was carried out using the QuikChangemutagenesis kit (Stra-tagene). The gene encoding BxGH99 was constructed from synthesizedoligonucleotide fragments (Genscipt, Inc.) and also subcloned into pET28a.Protein production and purification was identical for both Bt- and BxGH99enzymes, including all respective variants. Escherichia coli BL21 (DE3) cellsharboring the GH99-encoding plasmid were cultured in 0.5 L ZYM-5052 auto-induction media (39) supplemented with 50 μgmL−1 kanamycin at 37 °C for8 h, with induction occurring overnight at 16 °C. Cells were harvested andresuspended in 50 mM NaH2PO4, pH 8.0, 300 mM NaCl, and lysed by sonica-tion. Soluble lysate was applied to a NiSO4-charged 5 mL HiTrap chelatingcolumn (GE Healthcare), preequilibrated in the same buffer. The proteinwas eluted in an imidazole gradient, dialyzed, concentrated, and further pur-ified on an S75 16∕60 gel filtration column (GE) preequilibrated in 25 mMHepes, pH 8.0, 50 mM NaCl. The BtGH99 selenomethionine derivative wasoverexpressed in PASM-5052 media (39), otherwise all isolation and purifica-tion steps were as described above.

Activity, Kinetics and Stereochemistry. GH99 activity on GlcMan9GlcNAc2 wasstudied by MALDI-TOF mass spectrometry of the permethylated products fol-lowing overnight incubation at 37 °C (see SI Methods). The ligand affinity ofBtGH99 for Glc-DMJ and Glc-IFG was analyzed by ITC using an iTC200 calori-meter (MicroCal). Assays were carried out at 25 °C with Glc-DMJ (6.4 mM) andGlc-IFG (3.0 mM) titrated into the ITC cell containing 460 and 370 μM BtGH99,respectively. The dissociation constant for each reaction (Kd) was then calcu-lated using the Origin 7 software package (MicroCal). Kinetic parameters forthe hydrolysis of the synthetic substrate α-glucopyranosyl-1,3-α-mannopyra-nosyl fluoride (Glc-ManF) were determined using a fluoride-selective elec-trode with NMR analysis used to determine reaction stereochemistry (seeSI Methods).

Crystallization, Data Collection, and Structure Solution. The BtGH99 structurewas solved using single wavelength anomalous dispersion techniques usingthe selenomethionyl protein with data collected at beamline I24 of the Dia-mond Light Source. Other structures were solved by molecular replacementwith data collected on beamlines ID23-2 and ID14-1, respectively, of the Eur-opean Synchrotron Radiation Facility, and at beamlines I04-1 and I03 of theDiamond Light Source. Full details of crystallization, data collection, andstructure solution, including programs used, are given in the SI Methods.

ACKNOWLEDGMENTS. G.J.D. thanks the Biotechnology and Biological SciencesResearch Council for funding and is a Royal Society/Wolfson Research Meritaward recipient. T.M.G. is a Sir Henry Wellcome Fellowship recipient.S.J.W. thanks the Australian Research Council and the School of Chemistry,University of Melbourne, for funding support. T.W. thanks the NetherlandsOrganisation for Scientific Research for funding support. The York Center ofExcellence in Mass Spectrometry was created thanks to a major capital invest-ment through Science City York, supported by Yorkshire Forward with fundsfrom the Northern Way Initiative.

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