N-glycan degradation by bacterial glycoside phosphorylases 1 Role of glycoside-phosphorylases in mannose foraging by human gut bacteria*

In order to metabolise both dietary fibre constituent carbohydrates and host glycans lining the intestinal epithelium, gut bacteria produce a wide range of carbohydrate active enzymes, of which glycoside hydrolases are the main components. In this study, we describe the ability of phosphorylases to participate in the breakdown of human Nglycans, from an analysis of the substrate specificity of UhgbMP, a mannoside phosphorylase of the GH130 protein family discovered by functional metagenomics. UhgbMP is found to phosphorolyze β-DManp-1,4-β-D-GlcpNAc-1,4-D-GlcpNAc, and is also highly efficient enzyme to catalyze the synthesis of this precious N-glycan core oligosaccharide by reverse-phosphorolysis. Analysis of sequence conservation within family GH130, mapped on a 3D model of UhgbMP and supported by site-directed mutagenesis results, revealed two GH130 subfamilies, and allowed the identification of key residues responsible for catalysis and substrate specificity. The analysis of the genomic context of 65 known GH130 sequences belonging to human gut bacteria indicates that the enzymes of the GH130_1 subfamily would be involved in mannan catabolism, while the enzymes belonging to the GH130_2 subfamily, would rather work in synergy with glycoside hydrolases of the GH92 and GH18 families in the breakdown of N-glycans. The use of GH130 inhibitors as therapeutic agents or functional foods, could thus be considered as an innovative strategy to inhibit N-glycan degradation, with the ultimate goal of protecting, or restoring, the epithelial barrier. http://www.jbc.org/cgi/doi/10.1074/jbc.M113.483628 The latest version is at JBC Papers in Press. Published on September 16, 2013 as Manuscript M113.483628 Copyright 2013 by The American Society for Biochemistry and Molecular Biology, Inc. by gest on N ovem er 9, 2017 hp://w w w .jb.org/ D ow nladed from N-glycan degradation by bacterial glycoside phosphorylases 2 The human gut microbiota is a dense and complex ecosystem, which plays a crucial role in maintaining human health. The many meta-omic studies carried out in the last few years have shown that certain metabolic diseases, such as obesity and some inflammatory diseases, such as Inflammatory Bowel Diseases (IBD) [1– 4], are associated with structural and functional imbalances of this microbiota ([5] for review). The catabolism of complex carbohydrates by gut bacteria plays a key role in the microbial colonisation and equilibrium of the digestive tract [6,7]. Indeed, the plant polysaccharides (cellulose, hemicelluloses, pectin, and resistant starch) that compose dietary fibre supply the main source of carbon for gut bacteria, in the form of monosaccharides produced by polysaccharide breakdown, such as glucose, xylose, arabinose, uronic acids, and to a lesser extent, galactose, mannose and rhamnose [8]. In particular, mannose is found in plant cell walls, in mannan and gluco-mannan, the backbones of which respectively consisting of a homopolymer of β1,4-linked D-mannopyranosyl residues, and a heterogenous sequence of β1,4-linked Dglucopyranosyl and D-mannopyranosyl units [9]. There is however a further source of complex carbohydrates in the gut, namely the heavily Oand N-glycosylated glycoproteins that line the intestinal epithelium to form a protective barrier against pathogens, and chemical and mechanical aggression [10]. While host O-glycans are rich in N-acetyl-hexosamines (GlcNAc, GalNAc and Neu5Ac), galactose and fucose, many mature Nglycans contain 8 αand β-linked Dmannpyranosyl residues linked to chitobiose [11] (Figure 1). Human glycan–microbial interactions in the gastro-intestinal tract play a crucial role in determining the outcome of relations of both commensals and pathogens with the host. Indeed, alterations in the structure and/or quantity of host glycans due to biosynthetic defects or microbial degradation alter their barrier function and are thought to be involved in the initiation and the maintenance of mucosal inflammation in inflammatory bowel diseases (IBDs), and in the development of intestinal cancer [12]. In order to breakdown these complex carbohydrates of either plant or human origin, gut bacteria produce a full repertoire of Carbohydrate-active enzymes (CAZymes, listed in the CAZy database [13]), of which glycosidehydrolases (GHs) are the main constituents, as revealed by Gill et al [14]. In particular, the degradation of dietary mannans requires both endo-β-mannanases and β-mannosidases to release mannose [15]. The hydrolysis of Nglycans by a broad consortium of endoand exo, α-and β-glycosidases, in particular those produced by the prominent gut bacterium B. thetaiotaomicron [16–20] has also been described. Until August, 2013, only glycoside hydrolases (GHs) have been implicated in Nglycan breakdown. However, other types of CAZymes participate in the breakdown of complex carbohydrates, particularly those of plant origin, by working in synergy with GHs: carbohydrate esterases (CEs), polysaccharide lyases (PLs) and glycoside phosphorylases (GPs). The presence of CEs and PLs is readily detectable in gut bacterial genomes, metagenomes and metatranscriptomes, since they are sufficiently divergent from GHs as to be classified in their own CAZyme families. This is not the case for GPs, which are found both in the glycosyl transferase (GT) and glycoside hydrolase families, depending on the sequence and catalytic mechanism similarities shared with GT and GH archetypes, respectively. The prevalence of GPs and their role in the metabolism of carbohydrates is therefore difficult to evaluate on the basis of sequence data alone, as a given family of GHs (or of GTs) may comprise both GHs (or GTs) as well as GPs. GPs catalyse the breakdown of a glycosidic linkage from oligosaccharide or polysaccharide substrates with concomitant phosphate glycosylation, to yield a glycosyl-phosphate product and a sugar chain of reduced length. These enzymes are also able to perform reverse phosphorolysis (in the so called “synthetic reaction”) to form a glycosidic bond between the glycosyl unit originating from the glycosylphosphate, which acts as the sugar donor, and a carbohydrate acceptor [21–23]. Retaining GPs, for which phosphorolysis occurs via overall retention of the substrate anomeric configuration, are found in CAZy families GT4, GT35 and GH13 families. Inverting GPs are classified in families GH65, GH94, GH112 and GH130. Inverting GH-related GPs and hydrolytic enzymes use a similar single displacement mechanism, differing in the requirement of GPs for a single catalytic residue (the proton donor), and inverting GHs for two catalytic residues [22]. In GP-catalysed reactions, the reaction begins with the direct nucleophilic attack by phosphate to the glycosidic bond with the aid of by gest on N ovem er 9, 2017 hp://w w w .jb.org/ D ow nladed from N-glycan degradation by bacterial glycoside phosphorylases 3 the catalytic residue, which donates a proton to the glycosidic oxygen atom, and then proceeds through an oxocarbenium cation-like transition state. In GH reaction mechanisms, the nucleophilic attack of the C1 of the glycoside is performed by a water molecule activated by the catalytic base. The natural structural and functional diversity of GPs thus appears to be highly restricted, since (i) they are found in only seven of the 226 GH and GT families listed in the CAZy database (March 2013); (ii) approximately only 15 EC entries are currently assigned to GPs [24], (iii) their specificity towards glycosyl phosphates is limited to αand β-D-glucopyranose-1-phosphate [25–29] which are the most prevalent substrates, α-Dgalactopyranose-1-phosphate [30], N-acetyl-α-Dglucosamine-1-phosphate [31], and α-Dmannopyranose-1-phosphate [32]. In July 2013, α-D-mannopyranose-1-phosphate specificity was described for just one enzyme produced by a human gut bacterium, the B. fragilis NCTC 9343 mannosylglucose phosphorylase (BfMP), which converts β-D-mannopyranosyl-1,4-Dglucopyranose and phosphate into α-Dmannopyranose-1-phosphate and D-glucose [32]. This enzyme has been implicated in the catabolism of linear dietary mannans, assisted by a β-1,4-mannanase and a mannobiose 2epimerase. During review of the present manuscript, Nihira et al. [33] reported the discovery of a metabolic pathway for N-glycans, that includes a β-D-mannosyl-N-acetyl-1,4-Dglucosamine phosphorylase named BT1033, produced by the human gut inhabitant B. thetaiotamicron VPI-5482. BfMP and BT1033 are two of the four enzymes to be characterized in the recently created GH130 family, which includes a total of 447 entries, from archaea, bacteria, and eukaryotes. The other characterized GH130 enzymes are RaMP1 and RaMP2 from the ruminal bacterium Ruminococcus albus NE1 [34]. These enzymes have also been proposed to participate in mannan catabolism in the bovine rumen, and are assisted by an endo-mannanase and an epimerase, via RaMP2 and RaMP1catalysed phosphorolysis of β-1,4 mannooligosaccharides and 4-O-β-Dmannopyranosyl-D-glucopyranose, respectively [34]. X-ray crystallographic studies show that GH130 enzymes share a 5-fold β-propeller fold. Currently atomic coordinates data sets for four protein structures are available in the RCSB Protein Data Bank [35]: BACOVA_03624 protein from Bacteroides ovatus ATCC 8483 (3QC2), BDI_3141 protein from Parabacteroides distasonis ATCC 8503 (3TAW), BT_4094 protein from Bacteroides thetaiotamicron VPI-5482 (3R67), and TM1225 protein from Thermotoga maritima MSB8 (1VKD). However, no function has yet been attributed to these four proteins, thus limiting the understanding of structure-specificity relations for GH130 enzymes and the investigation of their catalytic mechanism. Recently, the sequence of another GH130 enzyme, that we refer to as UhgbMP (Unknown human gut bacterium Mannoside Phosphorylase, GenBank accession number ADD61463.1), was discovered by functional metagenomics of the human gut microbiota [36]. The 36.6 kbp metagenomic DNA fragment containing the UhgbMP encoding gene was taxonomically assigned to an as yet unidentified bacterium belonging to the genus Bacteroides. However, UhgbMP itself presents 99% protein sequence identity with the hypothetical protein BACSTE_03540 from Bacteroides stercoris ATCC 43183 (accession number EDS13361.1). Here, we present an integrative approach, based on analyses of UhgbMP substrate specificity, and of metagenomic and genomic data at the level of the entire human gut ecosystem, to reveal the role of UhgbMP and of 64 other GH130 enzymes produced by known gut bacteria in the breakdown of host and dietary mannose-containing glycans. In addition, we establish the molecular basis of GH130 enzyme catalysis, supported by the experimental results of rational engineering of UhgbMP and the analysis of its three-dimensional molecular model. Finally, we discuss the potential of this enzyme for the development of therapeutic agents and functional foods to inhibit N-glycan degradation, with the ultimate goal of protecting the epithelial barrier in the IBD context. EXPERIMENTAL PROCEDURES Recombinant UhgbMP production and purification First, the UhgbMP encoding gene was PCR amplified from the E. coli metagenomic clone (Genbank accession number GU942931) using primers forward 5’AGTATGAGTAGCAAAGTTATTATTCCTT GG 3’ and reverse 5’ TCAGATGATGCTTGTACGTTTGGTAAATT C 3’, by using the Expand Long Template PCR kit (Roche). by gest on N ovem er 9, 2017 hp://w w w .jb.org/ D ow nladed from N-glycan degradation by bacterial glycoside phosphorylases 4 To allow heterologous UhgbMP production in E. coli with His(6) tag at the Nterminal extremity, the PCR product was purified and subsequently cloned into the pCR8/GW/TOPO entry vector (Invitrogen), and then into the pDEST17 destination vector (Invitrogen), according to the manufacturer's recommendations. E.coli BL21-AI cells (Invitrogen) harboring the UhgbMP-encoding plasmid were cultured at 20°C for 24 hours in ZYM-5052 autoinduction medium [37] supplemented with 100 μg/mL ampicillin, inoculated at OD600nm 0.1. Cells were harvested and resuspended in 20mM Tris HCl, pH 7.0, 300 mM NaCl, and lysed by sonication. Soluble lysate was applied to a TALON resin loaded with cobalt (GE Healthcare) equilibrated in 20mM Tris HCl, pH 7.0, 300 mM NaCl. After column washing with 8 volumes of the same buffer supplemented with 10mM imidazole, the protein was eluted in 20mM Tris HCl, pH 7.0, 300 mM NaCl, 150 mM imidazole. Finally, the protein sample was desalted on a PD-10 column (GE Healthcare) and eluted in 20 mM Tris-HCl pH7.0, Tween 80 0.1% (vol/vol). In these conditions, 84 % of UhgbMP remained soluble after 8 days at 4°C, thus allowing further functional characterization. The purity of the purified wild-type UhgbMP and mutants was evaluated higher than 95 % by SDS page electrophoresis using Any kDTM MiniPROTEAN® TGXTM Precast Gel (BIO-RAD) (Supplementary Figure 1). After migration, proteins were stained with the PageBlue Protein Staining Solution (THERMO SCIENTIFIC) according to the manufacturer recommendations. Protein concentrations were determined by spectrometry using a NanoDrop® ND-1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA) The NanoDrop® measurement error was 5 %. The calculated extinction coefficient of the purified UhgbMP fused to an N-terminal His(6) tag is 76 630 M.cm. UhgbMP mutagenesis Site-directed mutagenesis was performed by using the pDEST17 plasmid harboring the wild type UhgbMP encoding gene as PCR template and the primers 5’ ATGGCTGTGCCAATACCGTAACCG 3’ and 5’ TACGGTATTGGCACAGCCATAGTAT3’ for D304N mutant, 5’ GTATGGCTACAACCCACGCGTGTGCT 3’ and 5’ ACGCGTGGGTTGTAGCCATACACCCA 3’ to obtain the D104N mutant, 5’ AACCGTACCAATGCATGGGCGATGT 3’ and 5’ ATGCATTGGTACGGTTCGCGC 3’ for the E273Q mutant, and finally 5’ TGGCGAGGACCCGCGCGT 3’ and 5’ TCCTCGCCATACACCCAGGTACCGA 3’ for the Y103E mutant. The PCR products, amplified with the Phusion® High-Fidelity DNA Polymerase (New England Biolabs), were purified and digested by DpnI (New England Biolabs) before E. coli TOP10 transformation. Protein production and purification were identical for wild-type UgbMP and its variants. Enzyme assays-All reactions were carried out at 37 °C (wild-type UhgbMP optimal temperature) in Tris HCl 20mM, pH 7.0 (wildtype UhgbMP optimal pH). Syntheses of mannooligosaccharides from α-D-mannopyranose-1phosphate were performed with 0.1 mg/ml purified UhgbMP during 24h at 37 °C in Tris HCl 20mM, pH 7.0, with 10 mM of α-Dmannopyranose-1-phosphate (Sigma, reference M1755), and 10 mM of D-glucose, D-mannose, D-galactose, D-fructose, N-acetyl-D-glucosamine, β-D-mannopyranosyl-1,4-N,N'-diacetyl chitobiose (Dextra, United Kingdom, reference MC0320), L-rhamnose, D-altrose, D-allose, Dfucose, L-fucose, D-mannitol, D-sorbitol, Dlyxose, xylitol, L-xylose, D-xylose, L-arabinose, or D-cellobiose. Phosphorolysis kinetic parameters were determined with 0.01 mg/ml of purified enzyme by quantifying α-D-mannose-1-phosphate release rate from 0.5 to 10 mM inorganic phosphate (10 mM corresponding to the intracellular concentration of inorganic phosphate that was previously measured in bacteria [38]) and 1 to 10 mM of β-D-mannopyranosyl-1,4-D-glucose (Carbosynth, United Kingdom, reference OM04754), 0.4 to 4 mM of β-1,4-D-mannan (Megazyme, Ireland, reference P-MANCB), 1 to 20 mM of β-D-mannopyranosyl-1,4-D-mannose (Megazyme, Ireland, reference O-MBI), 0.1 to 1 mM pNP-β-D-mannopyranose, or 0.05 to 0.5 mM of β-D-mannopyranosyl-1,4-N,N'-diacetyl chitobiose (Dextra, United Kingdom, reference MC0320). Reverse phosphorolysis kinetic parameters were determined with 0.01 mg/ml purified enzyme by quantifying α-Dmannopyranose-1-phosphate consumption rate from 0.5 to 10 mM α-D-mannopyranose-1phosphate and 5 to 40 mM of D-mannose, Dglucose, D-galactose, D-fructose, or N-acetyl-Dby gest on N ovem er 9, 2017 hp://w w w .jb.org/ D ow nladed from N-glycan degradation by bacterial glycoside phosphorylases 5 glucosamine, or 0.1 to 40 mM of N,N'-diacetyl chitobiose (Dextra, United Kingdom, reference C8002). The apparent kinetic parameters for phosphorolysis and reverse phosphorolysis at fixed initial concentrations of inorganic phosphate and α-D-mannose-1-phosphate, respectively, were determined by fitting the initial rates of α-D-mannose-1-phosphate release and consumption to the Michaelis-Menten equation. The kinetic parameters for phosphorolysis and synthesis of β-Dmannopyranosyl-1,4-N,N'-diacetyl chitobiose and β-D-mannopyranosyl-1,4-D-mannose were determined by varying carbohydrates, α-Dmannose-1-phosphate or inorganic phosphate concentrations, and by fitting the initial rates of α-D-mannose-1-phosphate release and consumption to the sequential random bi bi mechanism equation [39]. Non-linear regression was performed with SigmaPlot Enzyme Kinetics module, version 1.3 (Systat Software, Inc., San Jose California USA). UhgbMP specific activity towards β-1,4-D-manno-oligosaccharides, was determined with 0.1 mg/ml purified enzyme by quantifying α-D-mannopyranose-1-phosphate release rate from 10 mM inorganic phosphate and 10 mM of β-D-manno-oligosaccharides of polymerisation degree 2 to 6 (Megazyme, Ireland). The percentage of UhgbMP inhibition by exogeneous carbohydrates or polyols was measured with 0.1 mg/ml purified enzyme by quantifying α-D-mannopyranose-1-phosphate consumption rate from 10 mM α-Dmannopyranose-1-phosphate as glycosyl donor, with and without 10 mM of L-rhamnose, Daltrose, D-allose, D-fucose, D-mannitol, Dsorbitol, D-lyxose, xylitol, L-xylose, D-xylose, Larabinose, or D-cellobiose. The percentage of reversephosphorolysis activity of the D104N, D304N and E273Q variants, compared to that of the wild-type enzyme, was determined with 0.1 mg/ml of purified proteins by quantifying α-Dmannopyranose-1-phosphate consumption rate from 10 mM α-D-mannopyranose-1-phosphate as glycosyl donor and 10 mM D-mannose as acceptor. α-D-mannopyranose-1-phosphate was quantified by using high-performance anion exchange chromatography with pulsed amperometric detection (HPAEC-PAD). Carbohydrates and α-D-mannopyranose-1phosphate were separated on a 4 × 250 mm Dionex Carbopac PA100 column. A gradient of sodium acetate (from 0 to 150 mM in 15 min) and a isocratic step of 300 mM sodium acetate in 150 mM NaOH was applied at a 1 mL.min flow rate. Detection was performed using a Dionex ED40 module with a gold working electrode and a Ag/AgCl pH reference. Finally, the hydrolytic or phosphorolytic behavior of the wild-type UhgbMP and its Y103E variant was assessed by using 1 mM pNP-β-D-mannopyranose, in the absence, or presence, respectively, of 10 mM inorganic phosphate. The pNP release was monitored at OD405nm on a carry-100 UV-visible spectrophotometer (Agilent Technologies). Between 3 and 5 independent experiments were carried out to determine initial activity, kinetic constants, and percentage of inhibition by carbohydrates and polyols of wild-type UhgbMP and its mutants. For all reaction rate measurements, it was checked by HPAEC-PAD that less than 10 % of substrate was consumed, and that the amount of consumed or released αD-mannopyranose-1-phosphate increased linearly with time. NMR Spectroscopy-Freeze-dried reaction media were exchanged twice with 99.9 atom% D2O and lyophilized. Deuterium oxide was used as the solvent and sodium 2,2,3,3-tetradeuterio3-trimethylsilylpropanoate was selected as the internal standard. H and C NMR spectra were recorded on a Bruker Advance 500 MHz spectrometer using a 5 mm z-gradient TBI probe at 298 K, an acquisition frequency of 500.13 MHz and a spectral width of 8012.82 Hz. Spectra were acquired and processed using TopSpin 3.0 software. The various signals were assigned by comparison with signals obtained from α-D-mannopyranose-1-phosphate, β-Dmannopyranosyl-1,4-D-mannose (Megazyme, Irleland, reference O-MBI), β-Dmannopyranosyl-1,4-D-glucose (Carbosynth, United Kingdom, reference OM04754), or β-Dmannopyranosyl-1,4-N,N'-diacetyl chitobiose (Dextra, United Kingdom, reference MC0320), used as standards. Three-dimensional molecular modelling -The UhgbMP sequence was submitted to the ITASSER server for automated protein structure and function prediction [40]. The homologous Thermotoga maritima TM1225 structure (PDB accession code: 1VKD) was used to provide spatial restraints. The 3D model of UhgbMP predicted by I-TASSER was then further refined by energy minimisation using the CFF91 force by gest on N ovem er 9, 2017 hp://w w w .jb.org/ D ow nladed from N-glycan degradation by bacterial glycoside phosphorylases 6 field implementation in the DISCOVER module of the InsightII software suite (Accelrys, San Diego, CA, USA). The CFF91 cross terms, a harmonic bond potential, and a dielectric constant of 1.0 were specified in the energy function. An initial minimization was performed with positional restraints on the protein backbone using a steepest descent algorithm followed, by conjugated gradient minimization until the maximum RMS energy gradient was less than 0.5 kcal mol Å. The system was then fully relaxed without positional restraints. α-Dmannopyranose-1-phosphate, β-Dmannoheptaose and β-D-mannopyranosyl-1,4N,N'-diacetyl chitobiose were manually docked into the active site of UhgbMP. The ligand complexes were then optimized according to the minimization protocol described above with the ligand molecules free to move. Molecular graphics images were produced using PyMOL software (Schrödinger, LLC). GH130 multiple sequence alignment analyses -The 369 public sequences of GH130 enzymes listed in the CAZy database (www.cazy.org) in January 2013 were aligned with MUSCLE v3.7 [41]. A distance matrix was generated from the multiple sequence alignment using the BLOSUM62 amino acid residue substitution matrix. The output result file was subjected to hierarchical clustering using Ward’s method [39] and the resulting tree was visualized using DENDROSCOPE 3 [42]. Two sequence clusters were clearly apparent, members of which were accordingly assigned to the GH130_1 or GH130_2 sub-families. Position-dependent amino acid residue variation in multiple sequence alignment data was analysed using the Shannon information entropy measure (HX), calculated using SEQUESTER software. The Shannon entropy (HX) at residue alignment position (X), corrected for the normalized frequency of residue type occurrence, is computed as: