Structural and mechanistic insight into N-glycan processing by endo-α-mannosidase Andrew 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 a Department of Chemistry, University of York, Heslington, York YO10 5DD, United Kingdom; bSchool of Chemistry and Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, 30 Flemington Road, Parkville, Victoria 3010, Australia; cOxford Glycobiology Institute, Department of Biochemistry, 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
3D structure ∣ enzyme inhibition ∣ enzyme mechanism ∣ glycobiology ∣ glycosidase
N
-linked glycans are present on the majority of eukaryotic proteins and direct their folding and influence their stability. These polysaccharide decorations play important roles in processes such as protein folding, targeting, antigenicity, and lectin interactions, with defects leading to cellular dysfunction (1). Aberrant N-glycan composition, through either incorrect or incomplete processing, is associated with various conditions, including Alzheimer’s disease, congenital disorders of glycosylation, viral infection, and metastatic cancer progression (2–4). Alteration of N-glycan biosynthesis in cancerous cells and by human pathogenic viruses renders the various proteins involved in their biosynthesis and modification therapeutic targets. N-glycans are built-up and then “biosynthetically” degraded in an orchestrated process involving synthetic enzymes (glycosyltransferases) and catabolic enzymes (glycoside hydrolases) (5) (Fig. S1). Efforts to control diseases of glycoprotein biosynthesis have largely focused on the use of inhibitors of the “biosynthetic” trimming glycoside hydrolases involved in the early stages of N-glycan remodeling, such as glucosidases I and II. However, the failure of these inhibitors to provide an effective treatment of these conditions likely occurs, in part, because of a unique enzyme, endo-α-mannosidase, which cleaves mannoside linkages internally, within the first branch of an N-glycan chain (6–8). N-glycans are covalently bound to the side-chain nitrogen of asparagine (Asn) residues in the consensus Asn-Xxx-Ser/Thr (9). Within the endoplasmic reticulum (ER), presynthesized 14-mer www.pnas.org/cgi/doi/10.1073/pnas.1111482109
glycans are cotranslationally transferred en bloc by the multiprotein complex oligosaccharyltransferase from the glycophospholipid precursor, Glc3 Man9 GlcNAc2 -diphospho-dolichol, to Asn residues within nascent polypeptide chains (Fig. S1A). The initial processing steps of this 14-sugar glycan commence while the unfolded glycoprotein remains attached to the ribosome, and involves the sequential removal of the terminal α-1,2-glucose by glucosidase I and removal of both the subsequent and final α-1,3glucose moieties by glucosidase II, a procedure normally required for the formation of mature N-glycans (1, 5, 10) (Fig. S1B). Under normal conditions, the final α-1,3-glucose residue represents a checkpoint in protein folding and quality control because monoglucosylated immature N-glycans with a terminal α-Glc-1,3-αMan-1,2-α-Man chain are ligands for the molecular chaperones calnexin (CNX) and calreticulin (CRT) (11). The fate of newly synthesized glycoproteins is determined by a molecular inspection process that monitors their folding state. Correctly folded proteins exit the CNX/CRT cycle with removal of the final α1,3-glucose from the Man9 GlcNAc2 polysaccharide followed by α-mannosidase I cleavage of a mannose residue from the first branch and translocation to the Golgi apparatus. Misfolded proteins undergo cycles of deglucosylation and reglucosylation, catalyzed by luminal UDP-glucose-dependent glycoprotein glycosyltransferase. Terminally misfolded proteins are extracted from the folding cycle in a process termed ER-associated degradation and are retrotranslocated to the cytosol, where they are ubiquitinylated and proteasomally degraded. Endo-α-mannosidase (classified into Carbohydrate-Active Enzymes Database family GH99; refs. 12 and 13; www.cazy.org) provides a glucosidase I and II independent pathway for the maturation of N-glycans (14). Endo-α-mannosidase hydrolyzes the α-1,2-mannosidic bond between the glucose-substituted mannose and the remainder of the N-glycan, and acts on the structures Glc1–3 Man9 GlcNAc2 as well as structures that have been trimmed 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 subcelAuthor 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 the Protein Data Bank, www.pdb.org (PDB ID codes 4acy, 4acz, 4ad0, 4ad1, 4ad2, 4ad3, 4ad4, and 4ad5). 1
To whom correspondence may be addressed. E-mail:
[email protected] 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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CHEMISTRY
N-linked glycans play key roles in protein folding, stability, and function. Biosynthetic modification of N-linked glycans, within the endoplasmic reticulum, features sequential trimming and readornment steps. One unusual enzyme, endo-α-mannosidase, cleaves mannoside 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 and mechanistic dissection of endo-α-mannosidase. Structures solved at resolutions 1.7–2.1 Å reveal a ðβ∕αÞ8 barrel fold in which the catalytic center is present in a long substrate-binding groove, consistent with cleavage within the N-glycan chain. Enzymatic cleavage of authentic Glc1∕3 Man9 GlcNAc2 yields Glc1∕3 -Man. Using the bespoke substrate α-Glc-1,3-α-Man fluoride, the enzyme was shown to act with retention of anomeric configuration. Complexes with the established endo-α-mannosidase inhibitor α-Glc-1,3-deoxymannonojirimycin and a newly developed inhibitor, α-Glc-1,3isofagomine, and with the reducing-end product α-1,2-mannobiose structurally define the −2 to þ2 subsites of the enzyme. These structural and mechanistic data provide a foundation upon which to develop new enzyme inhibitors targeting the hijacking of N-glycan synthesis in viral disease and cancer.
BIOCHEMISTRY
Edited by Chi-Huey Wong, Academia Sinica, Taipei, Taiwan, and approved November 28, 2011 (received for review August 9, 2011)
lular localization of endo-α-mannosidase (14), no structures have been reported and nothing is known about its catalytic mechanism. Endo-α-mannosidase therefore represents an important enzyme for further study, with the ultimate goal of developing approaches to treat diseases involving aberrant N-glycosylation. We present a structural and kinetic analysis of two GH99 endoα-mannosidase enzymes from the enteric bacteria Bacteroides thetaiotaomicron (BtGH99) and Bacteroides xylanisolvens (BxGH99), the first structures for any GH99 enzyme. We show that these protein orthologs are endo-α-mannosidases active on glucosylated N-glycans and perform catalysis with a net retention of anomeric configuration. Structures of complexes obtained with two aza/imino sugar inhibitors reveal intimate details of the catalytic residues, allowing the proposal of a unique blueprint for catalysis and providing a structural rationale for inhibition in both a mechanistic context and ultimately for the development of therapeutic agents. Results Activity and Kinetics of GH99 Endo-α-Mannosidase. B. thetaiotaomi-
cron and B. xylanisolvens GH99 enzymes display 42% and 41% sequence identity, respectively, to Homo sapiens endo-α-mannosidase (sequence alignments in Fig. S2). In order to ascertain whether the Bacteroides enzymes were appropriate models of the human enzyme, activity on GlcMan9 GlcNAc2 was studied by mass spectrometry. Both BtGH99 and BxGH99 catalyze the removal of a disaccharide from GlcMan9 GlcNAc2 , with peaks observed for both reaction products, yet have no activity on the unglucosylated substrate Man9 GlcNAc2 , indicative of conversion to Man8 GlcNAc2 through release of a disaccharide, likely GlcMan (Fig. 1A). We next studied the BtGH99-catalyzed hydrolysis of a fluorescently labeled mammalian glycan, Glc3 Man7 GlcNAc2 by normal phase HPLC (see SI Methods). BtGH99 shows biologically relevant activity against this substrate with K m and kcat ∕K m values of 83 μM and 2.6 s−1 mM−1 , respectively (Fig. S3). These data are in good agreement with the K m value of 55 μM determined for the rat liver enzyme studied by Lubas and Spiro (16). Thus, BtGH99 possesses specific α-1,2-mannosidase activity when the −1 position mannoside is modified by one or more α-1,3-linked glucose residues. No activity was observed on a range of aryl α-mannosides. To further probe the activity of this enzyme, the activated substrate, α-gluco-
pyranosyl-1,3-α-mannopyranosyl fluoride (Glc-ManF), was synthesized (see SI Methods). Hydrolysis of Glc-ManF releases fluoride, which is monitored using a fluoride ion-selective electrode. BtGH99 shows high activity against Glc-ManF with K m and kcat ∕K m values of 0.48 mM and 9.9 s−1 mM−1 (Fig. 1B). Although fluoride detection is constrained to a relatively limited pH range, further observation of this reaction across the pH range 5.5–7.5 revealed the enzyme to be optimally active at approximately pH 7.0, consistent with another bacterial homolog from Shewanella amazonensis (17). That Glc-ManF acts as a substrate although various aryl mannosides do not, coupled with activity on GlcMan9 GlcNAc2 and Glc3 Man9 GlcNAc2 , confirms BtGH99 as an endo-α-mannosidase with a requirement for a minimal α-1,3-linked disaccharide substrate and hence with obligate binding for catalysis in a −2 subsite (for subsite nomenclature see ref. 18). Further evidence for subsite specificity and the requirement for a minimal occupancy of the −2 and −1 subsites was obtained through binding of two BtGH99 inhibitors. α-Glucopyranosyl-1,3deoxymannojirimycin (Glc-DMJ) (Fig. 1C) is in widespread use as a specific endo-α-mannosidase inhibitor and was developed by modification of the known exo-α-mannosidase inhibitor deoxymannojirimycin with an endo-α-mannosidase-targeting 1,3-linked α-glucosyl residue (19, 20). Inspired by the success of this approach, a new inhibitor, α-glucosyl-1,3-isofagomine (Glc-IFG) (Fig. 1C), was synthesized (SI Methods) from the broad-spectrum azasugar inhibitor isofagomine, by introduction of a 1,3-linked α-glucosyl residue to capitalize upon beneficial binding interactions in the −2 subsite. Isothermal titration calorimetry (ITC) revealed that BtGH99 binds both Glc-DMJ and Glc-IFG, with K d values of 24 μM and 625 nM, respectively (Fig. 2D). The inhibition data support the requirement for a 1,3-linked disaccharide occupying the −1 and −2 subsites, which is further supported by analysis of crystal structures of inhibitor-enzyme complexes (see below). Three-Dimensional Structures of BtGH99 and BxGH99. The crystal structure of a selenomethionine derivative of the B. thetaiotaomicron GH99 endo-α-mannosidase enzyme was solved at a resolution of 1.70 Å (Table S1). Crystal structures of the native BtGH99 and 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 GlcMan9 GlcNAc2 (m∕z 2,600.0); (Upper) without enzyme and (Lower) with BtGH99 to yield a Hex8 GlcNAc2 product consistent with removal of Glc α-1,3-Man (the enzyme has no activity on Man9 GlcNAc2 ). (B) Michaelis–Menten kinetics of BtGH99 on the bespoke substrate Glc-Man fluoride (shown in inset). (C) The endo-α-mannosidase inhibitors Glc-DMJ (synthesis in ref. 19) and Glc-IFG (synthesis in this work). (D) Isothermal titration calorimetry of Glc-IFG binding to BtGH99. 782 ∣
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Active Site Structure and Catalysis. The apo-BtGH99 crystal form presented a major obstacle to the observation of enzyme-ligand complexes, owing to a large loop region (residues 64–79) projecting into the active-center of another molecule of BtGH99 forming an artefactual crystallographic dimer. Despite considerable effort, attempts to obtain a form of BtGH99 amenable to complex formation proved unsuccessful. A second bacterial ortholog, B. xylanisolvens endo-α-mannosidase, was therefore selected for structural characterization and proved amenable to complex formation, allowing structural determination of binary complexes with Glc-DMJ and Glc-IFG, and ternary complexes with each inhibitor and the reducing-end product α-1,2-mannobiose (Fig. 3). Imino and aza sugars have proved instrumental reagents in the glycosidase field (23, 24) both for structural and mechanistic dissection and as templates for therapeutic agents. Deoxynojirimycin-type iminosugars possess a basic nitrogen in place of the endocyclic ring oxygen and are mimics of an oxocarbenium-ion bearing positive charge on the ring oxygen position. Isofagominetype azasugars possess a basic nitrogen at the pseudoanomeric position, and can be considered mimics of a glycosyl cation with charge located on the anomeric carbon. These compounds are likely to prove effective inhibitors and high-affinity ligands of the various GH99 endo-α-mannosidase orthologs. The complexes of BxGH99 with Glc-DMJ (25) and Glc-IFG unveil the ligand– protein interactions within the −1 and −2 subsites (18) (Fig. 3).
BIOCHEMISTRY
cular replacement at resolutions of 2.00 and 1.90 Å, respectively. Together, the different crystal forms and asymmetric unit contents yield 10 crystallographically independent “views” of the structure. The three-dimensional structures of these bacterial endo-α-mannosidase enzymes reveal a single-domain protein adopting a ðβ∕αÞ8 barrel fold with a central open cleft formed from extended loop regions linking the secondary structural motifs comprising the catalytic active site, Fig. 2A. These loop regions adopt different conformations in the various structures determined, and given the extended and branching nature of the N-glycan substrate(s), it appears that these large flexible loops are capable of forming stabilizing interactions with the substrate, positioning the scissile α-1,2-mannosidic bond so as to promote catalysis within the active site. Perhaps unsurprisingly given the unique enzymatic activity assigned to GH99, this loop-adorned barrel motif appears unique, with both bacterial GH99 orthologs exhibiting no significant structural and sequence similarity to glycoside hydrolases from other families. Analysis against known structural motifs using the Dali server (21) reveals weak secondary structure matches, the closest being endo-β-1,4-mannanase from Trichoderma reesei (Protein Data Bank ID code 1QNR, Z ¼ 15.9, rmsd 3.1 Å across 233 Cα positions, sequence identity ¼ 8%). The GH99 active center, located at the terminus of a solventaccessible channel near the center of the barrel fold, possesses a cluster of carboxylate side chains likely to play a role in mannosidic bond hydrolysis. In order to help assign potential catalytic function to these residues, a time-course of BtGH99-catalyzed hydrolysis of Glc-ManF was studied by 1 H NMR spectroscopy. Enzymatic hydrolysis of the α-linked C–F bond resulted in the rapid appearance of a product, shown by 2D NOESY analysis to be α-Glc-1,3-α-mannose, thus indicating that GH99-catalyzed substrate hydrolysis occurs with a net retention of anomeric configuration (Fig. S4). Over several hours, the initial product was observed to undergo mutarotation, isomerizing to an equilibrium mixture of both α- and β-anomers at the reducing-end mannose. Classical retaining glycoside hydrolases operate through a twostep, double displacement reaction in which a covalent intermediate is formed and then hydrolyzed, via oxocarbenium-ion-like transition states (glycosidase mechanisms recently reviewed in ref. 22). In most cases, this displacement reaction is achieved by an enzymatic nucleophile such as a carboxylate (aspartate or glutamate) or, as for certain sialidases, a tyrosine residue. In some 2-acetamidoglycosidases (including families GH18, 20, 25, 56, 84, and 85), the nucleophile is the 2-acetamido group of the substrate, which acts in a “neighboring group” participation reaction. For GH99, we initially assumed that the catalytic apparatus of the enzyme comprised a catalytic nucleophile acting to form the covalent glycosyl-enzyme intermediate and a catalytic acid/base, first functioning as a general acid to assist leaving group departure and then as a general base to activate water for nucleophilic
CHEMISTRY
Fig. 2. Three-dimensional structure and conservation in GH99 endo-αmannosidases. BxGH99 in complex with Glc-IFG and α-1,2-mannobiose as a ribbon (A) and (B) a surface representation colored by sequence conservation (40) using the partial GH99 alignment as shown in Fig. S2.
attack at the anomeric carbon of the intermediate. Kinetic analysis of three BtGH99 active-center variants, E154A, E329A, and E332A (E156, E333, and E336 in BxGH99), reveals substantial decreases in catalytic activity (Table S2). Both the E154A and E329A variants show near zero activity. E332A meanwhile shows reduced activity with observed rate constants indicating an approximate 50-fold decrease in catalytic activity with respect to wild type using the activated Glc-ManF substrate. Against the natural substrate, Glc3 Man7 GlcNAc2 , the same mutant shows zero activity compared to wild type, under matching experimental conditions. In order to gain insight into the functions of the different components of the catalytic apparatus, 3D structures were determined with a bacterial endo-α-mannosidase in complex with inhibitors and the truncated reducing-end product, α-1,2mannobiose.
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-1,2mannobiose (in divergent stereo). Figures shown are REFMAC maximumlikelihood/σ A weighted 2F o − F c syntheses contoured between 0.26 and 0.32 electrons per Å3 . PNAS ∣
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Both inhibitors bind to the enzyme active site with the −1 ring located deep within a substrate-binding pocket in an undistorted 4 C conformation. The −2 glucosyl residue projects from the 1 catalytic cavity and appears solvent accessible. These subsites lie in the center of a substrate-binding cleft with a length of approximately 40 Å. The solvent-exposed nature of the −2 subsite glucoside allows the accommodation of elongated substrates with additional glucose residues at the 3 position. Aromatic platforms are common binding elements of carbohydrate active enzymes (26) and in these distal negative subsites are provided by a pair of highly conserved tryptophans at positions 48 and 126. Sequence alignments reveal a high degree of sequence conservation across various species in the region comprising the substrate-binding cleft (Fig. 2B). Indeed, all the interactions in the −2 to þ2 subsites appear to be essentially invariant (Figs. S2 (alignment) and S5 (interactions)). That the glucoside moiety interacts with residues in the −2 subsite, and also with the putative catalytic apparatus, provides a structural rationale for the catalytic requirement of a sugar occupying the −2 subsite of endo-α-mannosidase. Aza and imino sugar inhibitors have been used to identify the catalytic apparatus of glycosidases by virtue of the interactions of potential nucleophiles with the protonated nitrogen atoms (25, 26). BxGH99 structures in complex with Glc-DMJ and Glc-IFG reveal a water molecule, coordinated by Glu336 (332 in BtGH99) and poised below the pseudoanomeric nitrogen of Glc-IFG and the C1 of the DMJ moiety of Glc-DMJ, in the ideal position for nucleophilic attack to complete the second hydrolysis step of a classical double displacement retention mechanism. Glu336 is situated on the “α-face” of the sugar occupying the −1 subsite, in a position where it might interact with the glycosidic oxygen, thus fulfilling a role strongly indicative of the catalytic acid/base in the enzyme mechanism. Consistent with such a function, the BtGH99 variant E332A retains approximately 2% activity, compared to WT, on the activated fluoride substrate but has no detectable activity on the natural GlcMan9 GlcNAc2 glycan (determined by mass spectrometry) or against Glc3 Man7 GlcNAc2 (determined by HPLC).
A
Isofagomine-type azasugar inhibitors are useful crystallographic probes that typically allow direct observation of the enzymatic nucleophile of retaining glycoside hydrolases. Upon protonation, the pseudoanomeric nitrogen forms a salt bridge with the negatively charged nucleophile, acting as an “ionic trap” for the nucleophile, which during the reaction coordinate must approach within bonding distance of the anomeric carbon (23). All X-ray structures of retaining glycoside hydrolases with isofagomine-type compounds have, without exception, revealed the catalytic nucleophile to be located within 3 Å of the azasugar nitrogen, whereas for inverting enzymes, the nucleophilic water is found less than 3 Å away from nitrogen (Table S3). Of particular interest is the absence, in the GH99 orthologs, of any enzymatic carboxylate at a position suitable to act as the enzymatic nucleophile in a conventional double displacement reaction. Indeed, without invoking a conformational change, or repositioning of the −1 sugar residue, the BxGH99 complex structures show no protein residue in a position to act as a potential nucleophile; the most closely located candidates for a classical double displacement mechanism (shown in Fig. 4A) include the OE1 atom of Glu333 (approximately 3.5 Å distant) or the OH of Tyr46 or Tyr252 (4.0 Å distant). However, none of these residues appear close enough to interact with the inhibitor nitrogens at either ring position in the complexes (Movie S1). Furthermore, the observation of identical (and static) arrangements of the active site residues in three crystal forms of two enzymes with a total of 10 different packing arrangements provides no evidence of intrinsic conformational flexibility within the active site. Provocatively, it is well known that the alkaline solvolysis of α-mannosyl fluoride proceeds through a mechanism involving neighboring group participation in which the 2-OH attacks the anomeric position resulting in the formation of the intermediate 1,2-anhydro-βmannopyranose. This intermediate is opened by a nucleophile to afford an α-configured product possessing a retained anomeric configuration (27). Although no evidence currently supports the existence of a 1,2-anhydro sugar as an intermediate for glycosidase-catalyzed hydrolysis, it is possible that such a mechanism may occur with endo-α-mannosidase (Fig. 4B). Accordingly, the
enzyme HO
OH O
HO O HO HO HO HO
Nu
O
ROH
H
HO HO HO
Glu336
O
O O
O
O
O
O
O
H
H
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H
Glu336 O
HO HO HO
O
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O O
HO O HO OR
H
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HO O
HO O HO
O
Glu336
Glu333
H O
Nu
OH O
O
HO
HO
HO HO HO
Glu333
O
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OH O
HO O HO
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HO HO
H
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B
HO
OH enzyme O Nu
HO O HO OR
O
enzyme HO
O
H
HO HO
O H
Glu336 O
O O
HO O HO
HO
OH O
O
H
Glu336 O
Fig. 4. Putative catalytic mechanisms for GH99 endo-α-mannosidase. (A) Classical two-step double displacement mechanism proceeding via a glycosyl-enzyme intermediate; in the case of the structures reported here, this would require conformational change. (B) Neighboring group participation via a 1,2-anhydro sugar intermediate.
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Discussion Endo-α-mannosidase is a resident of the cis/medial compartment of the Golgi apparatus and pre-Golgi intermediates, and is a membrane-associated protein that is difficult to recombinantly express in soluble form (29). The bacterial orthologs from B. thetaiotaomicron and B. xylanisolvens are soluble proteins and may have been acquired by horizontal gene transfer because these organisms are common and beneficial components of the human gut (30). It has been suggested that, under normal conditions, endo-α-mannosidase acts to deglucosylate folded Glc1 Man7–9 glycoproteins that may reach the Golgi apparatus through being poor substrates for ER α-glucosidase II (7). The biological role of the bacterial enzymes is unclear, but may include, as is the case for the exoα-mannosidases of family GH92 (31), mannose foraging for basic metabolic inputs. Just as incomplete deglucosylation in the mammalian cell prevents the action of exo-α-mannosidases, a similar problem with the bacterial exo-α-mannosidases might have led to the beneficial acquisition of the enzyme by the bacteria. It is notable that whereas the genomes of B. thetaiotaomicron and B. xylanisolvens possess many copies of the N-glycan active exo-αmannosidases (24 and 17 GH92 enzymes, respectively; 3 GH125 enzymes each) each genome contains only a single copy of a GH99 enzyme, which supports its role in relieving a metabolic bottleneck in the degradation of glucosylated N-glycans. Alternatively, or additionally, the bacterial endo-α-mannosidase may act on yeast mannans, which contain the epimeric α-mannosyl-1,3-mannose substructure. In this regard, it is interesting that studies of mammalian endo-α-mannosidase have indeed shown that α-mannosyl-
1,3-deoxymannonojirimycin, the epimer of Glc-DMJ, is an effective inhibitor (20). Mammalian endo-α-mannosidase has a preference for monoglucosylated N-glycans and its activity increases as the number of mannose residues on the 6′-mannose branch decreases (16). The structures of the bacterial orthologs suggest that the majority of substrate specificity is for the −1 and −2 subsites with additional sugars off the 3-position of the −2 glucose residue likely to project into solvent and not to be involved in significant enzyme interactions. α-Mannosidases must overcome certain substrate-specific challenges to achieve effective catalysis because the enhanced anomeric effect in mannose deters substrate distortion, and thus an attacking nucleophile suffers destabilizing 1,2diaxial interactions with the axial 2-OH group. In retaining GH38 (32), and inverting GH47 (33) GH92 (31) α-mannosidases, a divalent metal ion (Zn2þ for GH38, Ca2þ for GH47 and GH92) is found to coordinate the 2- and 3-hydroxyls, possibly assisting in catalysis through substrate distortion and in delivery of the nucleophile (31). By contrast, in the inverting GH125 enzymes no metal ion is observed in any structures and no substrate distortion was seen in a pseudo-Michaelis complex with an S-linked disaccharide spanning the active site (34). It is unclear how this family overcomes the destabilizing diaxial interaction of the attacking water nucleophile, although it is possible that a favorable hydrogen bond between the 2-OH of the sugar and the nucleophilic water may assist in nucleophilic attack in a similar way to the divalent metals in the GH38, GH47, and GH92 enzymes. A notable feature of the GH99 active site observed in both bacterial endo-α-mannosidase orthologs is the absence of a metal ion or indeed of a site for possible metal ion coordination. The enzyme-inhibitor complexes with Glc-DMJ and Glc-IFG determined 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 conserved residue, Glu333, is found in close proximity to O2, and its mutagenesis results in a near-complete decrease in catalytic activity. Taken together, these observations suggest that the hydrolysis reaction catalyzed by the GH99 endo-α-mannosidase family proceeds via both a metal-independent and possibly a uniquely unorthodox mechanism. Inhibitors that intervene in early steps of N-glycan biosynthesis have been largely ineffective in suppressing N-linked glycan processing. Cells in which glucosidase II has been inhibited or subjected to genetic knockout retain 40–80% normal N-glycan processing function through the intervention of endo-α-mannosidase (8, 35, 36). During a castanospermine (CST)-imposed α-glucosidase I and II blockade, HepG2 cells produced N-linked glycoproteins with the normal glycan structure and resulted in concomitant release of free Glc1–3 Man oligosaccharides (6). Evidence for the basal action of endo-α-mannosidase in the absence of glucosidase inhibition or knockout was demonstrated through the use of the mannosidase I inhibitor, DMJ, which prevents further processing of the deglucosylated N-linked oligosaccharides. Man6–8 GlcNAc, but not Man9 GlcNAc, structures were identified, with the Man8 GlcNAc structure being the characteristic isomer generated by endo-α-mannosidase cleavage (6). Coadministration of the α-glucosidase I and II inhibitor CST and the selective endo-α-mannosidase inhibitor Glc-DMJ resulted in inhibition of biosynthesis of Glc3 Man and N-linked glycoproteins (25). On the other hand, Golgi α-mannosidase II is seen as a promising target for therapeutic intervention, as this enzyme acts after endo-α-mannosidase rejoins the normal pathway and so acts on all N-linked glycans undergoing processing (25). Swainsonine, a selective inhibitor of Golgi α-mannosidase II, blocks the abnormal formation of complex oligosaccharides, leading to reduced metastasis and tumor growth (37). However, swainsonine also inhibits the closely related lysosomal α-mannosidase, limiting its clinical use (38). Combination therapies of glucosidase I/II inhi-
CHEMISTRY
kinetically essential and conserved active-center residue Glu333, poised 2.6 Å away from the O2 group of the substrate, would deprotonate the 2-OH, promoting its nucleophilic attack on C1. The anti-arrangement of O2 and the leaving group seen in the complex with Glc-DMJ is that stereoelectronically required for epoxide formation, which requires a linear transition state of the attacking nucleophile, the reactive carbon center, and the leaving group. Neighboring group participation by the 2-acetamido group of N-acetyl-β-glycosides is now well established in families GH18, 20, 25, 56, 84, and 85 (28). Together the precedent for neighboring group participation in these glycoside hydrolase families, the lack of an apparent enzymatic nucleophile in the GH99 structures, and the established neighboring group participation mechanism for nonenzymatic hydrolysis of α-mannosyl fluoride, provides an enticing precedent to suggest this unorthodox enzymatic mechanism. In order to dissect the leaving-group (“positive” numbers; ref. 18) subsites of the GH99 endo-α-mannosidase, additional crystallization experiments were conducted with either Glc-DMJ or Glc-IFG together with α-1,2-mannobiose. Such conditions afford complexes with the inhibitors in the −1 and −2 subsites, and with the mannobiose disaccharide in the þ1 and þ2 subsites. As with the −2 moiety, the þ2 mannose residue appears solvent accessible, projecting outside the active site cavity at a distance of approximately 7 Å from α-1,3-glucose, consistent with an ability to accommodate the remaining reducing end residues of a complex N-glycan. Phenylalanines at positions 253 and 258 provide an aromatic platform positioning the þ1 mannoside, with Tyr289 and Asn298 making hydrogen-bonding interactions with the O3 and O4 positions of the same sugar. Likewise within the þ2 subsite, hydrogen bonding occurs between O3 and O4 and Tyr195, Asp196, and Tyr198. Despite the sugar residue occupying the þ1 subsite appearing to be slightly displaced from its likely true position during hydrolytic cleavage of the authentic substrate (O2 of the þ1 sugar lies approximately 4 Å from the atom equivalent to the anomeric C1 of the residue in the −1 subsite in both inhibitor complexes), both structures show the carboxylate group of the likely catalytic acid/base, Glu336, to be in close proximity to the position of the scissile α-1,2-mannosidic bond.
bitors (e.g., 6-O-butanoyl-CST) and endo-α-mannosidase inhibitors may provide a viable alternative approach. The development of Glc-IFG as a more potent inhibitor of the bacterial endo-αmannosidase serves as a valuable precedent for glucosylation of inhibitors of exo-α-mannosidases to search for more active inhibitors of endo-α-mannosidase, which will be assisted by the structural and mechanistic blueprint provided by this work. Methods Gene Cloning, Mutagenesis and Protein Production. The gene encoding BtGH99 was amplified from genomic DNA by PCR and ligated into a modified pET28a vector encoding an N-terminal His6 -tag (pETLIC-BtGH99). Site-directed mutagenesis was carried out using the QuikChange mutagenesis kit (Stratagene). The gene encoding BxGH99 was constructed from synthesized oligonucleotide fragments (Genscipt, Inc.) and also subcloned into pET28a. Protein production and purification was identical for both Bt- and BxGH99 enzymes, including all respective variants. Escherichia coli BL21 (DE3) cells harboring the GH99-encoding plasmid were cultured in 0.5 L ZYM-5052 autoinduction media (39) supplemented with 50 μg mL−1 kanamycin at 37 °C for 8 h, with induction occurring overnight at 16 °C. Cells were harvested and resuspended in 50 mM NaH2 PO4 , pH 8.0, 300 mM NaCl, and lysed by sonication. Soluble lysate was applied to a NiSO4 -charged 5 mL HiTrap chelating column (GE Healthcare), preequilibrated in the same buffer. The protein was eluted in an imidazole gradient, dialyzed, concentrated, and further purified on an S75 16∕60 gel filtration column (GE) preequilibrated in 25 mM Hepes, pH 8.0, 50 mM NaCl. The BtGH99 selenomethionine derivative was overexpressed in PASM-5052 media (39), otherwise all isolation and purification steps were as described above. 1. Molinari M (2007) N-glycan structure dictates extension of protein folding or onset of disposal. Nat Chem Biol 3:313–320. 2. Akasaka-Manya K, et al. (2010) Protective effect of N-glycan bisecting GlcNAc residues on beta-amyloid production in Alzheimer’s disease. Glycobiology 20:99–106. 3. Damme M, et al. (2010) Impaired lysosomal trimming of N-linked oligosaccharides leads to hyperglycosylation of native lysosomal proteins in mice with alpha-mannosidosis. Mol Cell Biol 30:273–283. 4. Zhao YY, et al. (2008) Functional roles of N-glycans in cell signaling and cell adhesion in cancer. Cancer Sci 99:1304–1310. 5. Herscovics A (1999) Importance of glycosidases in mammalian glycoprotein biosynthesis. Biochim Biophys Acta 1473:96–107. 6. Moore SE, Spiro RG (1990) Demonstration that Golgi endo-alpha-D-mannosidase provides a glucosidase-independent pathway for the formation of complex N-linked oligosaccharides of glycoproteins. J Biol Chem 265:13104–13112. 7. Moore SE, Spiro RG (1992) Characterization of the endomannosidase pathway for the processing of N-linked oligosaccharides in glucosidase II-deficient and parent mouse lymphoma cells. J Biol Chem 267:8443–8451. 8. Rabouille C, Spiro RG (1992) Nonselective utilization of the endomannosidase pathway for processing glycoproteins by human hepatoma (HepG2) cells. J Biol Chem 267:11573–11578. 9. Kornfield R, Kornfield S (1985) Assembly of asparagine-linked oligosaccharides. Annu Rev Biochem 54:631–664. 10. Ruddock LW, Molinari M (2006) N-glycan processing in ER quality control. J Cell Sci 119:4373–4380. 11. Helenius A, Aebi M (2004) Roles of N-linked glycans in the endoplasmic reticulum. Annu Rev Biochem 73:1019–1049. 12. Henrissat B, Davies G (1997) Structural and sequence-based classification of glycoside hydrolases. Curr Opin Struct Biol 7:637–644. 13. Henrissat B (1991) A classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem J 280:309–316. 14. Roth J, et al. (2003) The role of glucosidase II and endomannosidase in glucose trimming of asparagine-linked oligosaccharides. Biochimie 85:287–294. 15. Dairaku K, Spiro RG (1997) Phylogenetic survey of endomannosidase indicates late evolutionary appearance of this N-linked oligosaccharide processing enzyme. Glycobiology 7:579–586. 16. Lubas WA, Spiro RG (1988) Evaluation of the role of rat liver Golgi endo-alpha-D-mannosidase in processing N-linked oligosaccharides. J Biol Chem 263:3990–3998. 17. Matsuda K, et al. (2011) Heterologous Expression, Purification, and Characterization of an alpha-Mannosidase Belonging to Glycoside Hydrolase Family 99 of Shewanella amazonensis. Biosci Biotechnol Biochem 75:797–799. 18. Davies GJ, Wilson KS, Henrissat B (1997) Nomenclature for sugar-binding subsites in glycosyl hydrolases. Biochem J 321:557–559. 19. Spreitz J, Stutz AE (2004) Golgi endomannosidase inhibitor, alpha-D-glucopyranosyl(1 → 3)-1-deoxymannojirimycin: A five-step synthesis from maltulose and examples of N-modified derivatives. Carbohydr Res 339:1823–1827. 20. Ardron H, et al. (1993) Synthesis of 1,5-dideoxy-3-O-(α-D-mannopyranosyl)-1,5-iminoD-mannitol and 1,5-dideoxy-3-O-(α-D-glucopyranosyl)-1,5-imino-D-mannitol—powerful inhibitors of endomannosidase. Tetrahedron Asymmetry 4:2011–2024.
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Activity, Kinetics and Stereochemistry. GH99 activity on GlcMan9 GlcNAc2 was studied by MALDI-TOF mass spectrometry of the permethylated products following overnight incubation at 37 °C (see SI Methods). The ligand affinity of BtGH99 for Glc-DMJ and Glc-IFG was analyzed by ITC using an iTC200 calorimeter (MicroCal). Assays were carried out at 25 °C with Glc-DMJ (6.4 mM) and Glc-IFG (3.0 mM) titrated into the ITC cell containing 460 and 370 μM BtGH99, respectively. The dissociation constant for each reaction (K d ) was then calculated using the Origin 7 software package (MicroCal). Kinetic parameters for the hydrolysis of the synthetic substrate α-glucopyranosyl-1,3-α-mannopyranosyl fluoride (Glc-ManF) were determined using a fluoride-selective electrode with NMR analysis used to determine reaction stereochemistry (see SI Methods). Crystallization, Data Collection, and Structure Solution. The BtGH99 structure was solved using single wavelength anomalous dispersion techniques using the selenomethionyl protein with data collected at beamline I24 of the Diamond Light Source. Other structures were solved by molecular replacement with data collected on beamlines ID23-2 and ID14-1, respectively, of the European Synchrotron Radiation Facility, and at beamlines I04-1 and I03 of the Diamond Light Source. Full details of crystallization, data collection, and structure solution, including programs used, are given in the SI Methods. ACKNOWLEDGMENTS. G.J.D. thanks the Biotechnology and Biological Sciences Research Council for funding and is a Royal Society/Wolfson Research Merit award 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 Netherlands Organisation for Scientific Research for funding support. The York Center of Excellence in Mass Spectrometry was created thanks to a major capital investment through Science City York, supported by Yorkshire Forward with funds from the Northern Way Initiative. 21. Holm L, Rosenström P (2010) Dali server: Conservation mapping in 3D. Nucleic Acids Res 38(Suppl 2):W545–W549. 22. Vocadlo DJ, Davies GJ (2008) Mechanistic insights into glycosidase chemistry. Curr Opin Chem Biol 12:539–555. 23. Lillelund VH, Jensen HH, Liang X, Bols M (2002) Recent developments of transitionstate analogue glycosidase inhibitors of non-natural product origin. Chem Rev 102:515–554. 24. Gloster TM, Davies GJ (2010) Glycosidase inhibition: Assessing mimicry of the transition state. Org Biomol Chem 8:305–320. 25. Hiraizumi S, Spohr U, Spiro RG (1993) Characterization of endomannosidase inhibitors and evaluation of their effect on N-linked oligosaccharide processing during glycoprotein biosynthesis. J Biol Chem 268:9927–9935. 26. Nerinckx W, Desmet T, Claeyssens M (2003) A hydrophobic platform as a mechanistically relevant transition state stabilising factor appears to be present in the active centre of all glycoside hydrolases. FEBS Lett 538:1–7. 27. Micheel F, Borrmann D (1960) A new method for the synthesis of saccharides larger cache (Translated from German). Chem Ber 93:1143–1147. 28. He Y, Macauley MS, Stubbs KA, Vocadlo DJ, Davies GJ (2010) Visualizing the reaction coordinate of an O-GlcNAc hydrolase. J Am Chem Soc 132:1807–1809. 29. Zuber C, Spiro MJ, Guhl B, Spiro RG, Roth J (2000) Golgi apparatus immunolocalization of endomannosidase suggests post-endoplasmic reticulum glucose trimming: Implications for quality control. Mol Biol Cell 11:4227–4240. 30. Qin J, et al. (2010) A human gut microbial gene catalogue established by metagenomic sequencing. Nature 464:59–65. 31. Zhu Y, et al. (2010) Mechanistic insights into a calcium-dependent family of α-mannosidases in a human gut symbiont. Nat Chem Biol 6:125–132. 32. Numao S, Kuntz DA, Withers SG, Rose DR (2003) Insights into the mechanism of Drosophila melanogaster Golgi α-Mannosidase II through the structural analysis of covalent reaction intermediates. J Biol Chem 278:48074–48083. 33. Vallée F, Karaveg K, Herscovics A, Moremen KW, Howell PL (2000) Structural basis for catalysis and inhibition of N-glycan processing class I α1,2-Mannosidases. J Biol Chem 275:41287–41298. 34. Gregg KJ, et al. (2011) Analysis of a new family of widely distributed metal-independent α-Mannosidases provides unique insight into the processing of N-linked glycans. J Biol Chem 286:15586–15596. 35. Fujimoto K, Kornfeld R (1991) alpha-Glucosidase II-deficient cells use endo alphamannosidase as a bypass route for N-linked oligosaccharide processing. J Biol Chem 266:3571–3578. 36. Volker C, et al. (2002) Processing of N-linked carbohydrate chains in a patient with glucosidase I deficiency (CDG type IIb). Glycobiology 12:473–483. 37. Goss PE, Baker MA, Carver JP, Dennis JW (1995) Inhibitors of carbohydrate processing: A new class of anticancer agents. Clin Cancer Res 1:935–944. 38. Colegate SM, Dorling PR, Huxtable CR (1979) A spectroscopic investigation of swainsonine: an α-mannosidase inhibitor isolated from Swainsona canescans. Aust J Chem 32:2257–2264. 39. Studier FW, Studier FW (2005) Protein production by auto-induction in high density shaking cultures. Protein Exp Purif 41:207–234. 40. Rockwell N, Lagarias JC (2007) Flexible mapping of homology onto structure with Homolmapper. BMC Bioinformatics 8(1):123–135.
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