Abstract
The metabolism of carbohydrate polymers drives microbial diversity in the human gut microbiota. It is unclear, however, whether bacterial consortia or single organisms are required to depolymerize highly complex glycans. Here we show that the gut bacterium Bacteroides thetaiotaomicron uses the most structurally complex glycan known: the plant pectic polysaccharide rhamnogalacturonan-II, cleaving all but 1 of its 21 distinct glycosidic linkages. The deconstruction of rhamnogalacturonan-II side chains and backbone are coordinated to overcome steric constraints, and the degradation involves previously undiscovered enzyme families and catalytic activities. The degradation system informs revision of the current structural model of rhamnogalacturonan-II and highlights how individual gut bacteria orchestrate manifold enzymes to metabolize the most challenging glycan in the human diet.
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Change history
05 April 2017
The PDB code ‘5MQP’ was added to the Data Availability section; and reference citations were corrected in the final paragraph of the main text.
References
Wißing, C. et al. Isotopic evidence for dietary ecology of late Neandertals in North-Western Europe. Quat. Int. 411, 327–345 (2016)
Pellerin, P. et al. Structural characterization of red wine rhamnogalacturonan II. Carbohydr. Res. 290, 183–197 (1996)
Apolinar-Valiente, R. et al. Polysaccharide composition of Monastrell red wines from four different Spanish terroirs: effect of wine-making techniques. J. Agric. Food Chem. 61, 2538–2547 (2013)
Cuskin, F. et al. Human gut Bacteroidetes can utilize yeast mannan through a selfish mechanism. Nature 517, 165–169 (2015)
Larsbrink, J. et al. A discrete genetic locus confers xyloglucan metabolism in select human gut Bacteroidetes. Nature 506, 498–502 (2014)
Rogowski, A. et al. Glycan complexity dictates microbial resource allocation in the large intestine. Nat. Commun. 6, 7481 (2015)
Koropatkin, N. M., Cameron, E. A. & Martens, E. C. How glycan metabolism shapes the human gut microbiota. Nat. Rev. Microbiol. 10, 323–335 (2012)
Martens, E. C. et al. Recognition and degradation of plant cell wall polysaccharides by two human gut symbionts. PLoS Biol. 9, e1001221 (2011)
Amaya, M. F. et al. Structural insights into the catalytic mechanism of Trypanosoma cruzi trans-sialidase. Structure 12, 775–784 (2004)
Lombard, V., Golaconda Ramulu, H., Drula, E., Coutinho, P. M. & Henrissat, B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 42, D490–D495 (2014)
Davis, B. G. et al. Tetrazoles of manno- and rhamno-pyranoses: Contrasting inhibition of mannosidases by 4.3.0 but of rhamnosidase by 3.3.0 bicyclic tetrazoles. Tetrahedron 55, 4489–4500 (1999)
Speciale, G., Thompson, A. J., Davies, G. J. & Williams, S. J. Dissecting conformational contributions to glycosidase catalysis and inhibition. Curr. Opin. Struct. Biol. 28, 1–13 (2014)
Fujita, K. et al. Molecular cloning and characterization of a β-l-Arabinobiosidase in Bifidobacterium longum that belongs to a novel glycoside hydrolase family. J. Biol. Chem. 286, 5143–5150 (2011)
Spellman, M. W., McNeil, M., Darvill, A. G., Albersheim, P. & Henrick, K. Isolation and characterization of 3-C-carboxy-5-deoxy-l-xylose, a naturally occurring, branched-chain, acidic monosaccharide. Carbohydr. Res. 122, 115–129 (1983)
Guérardel, Y. et al. The nematode Caenorhabditis elegans synthesizes unusual O-linked glycans: identification of glucose-substituted mucin-type O-glycans and short chondroitin-like oligosaccharides. Biochem. J. 357, 167–182 (2001)
Russa, R., Urbanik-Sypniewska, T., Choma, A. & Mayer, H. Identification of 3-deoxy-lyxo-2-heptulosaric acid in the core region of lipopolysaccharides from Rhizobiaceae. FEMS Microbiol. Lett. 84, 337–343 (1991)
Nagae, M. et al. Structural basis of the catalytic reaction mechanism of novel 1,2-α-l-fucosidase from Bifidobacterium bifidum. J. Biol. Chem. 282, 18497–18509 (2007)
Sulzenbacher, G. et al. Crystal structure of Thermotoga maritima α-l-fucosidase. Insights into the catalytic mechanism and the molecular basis for fucosidosis. J. Biol. Chem. 279, 13119–13128 (2004)
O’Neill, M. A., Ishii, T., Albersheim, P. & Darvill, A. G. Rhamnogalacturonan II: structure and function of a borate cross-linked cell wall pectic polysaccharide. Annu. Rev. Plant Biol. 55, 109–139 (2004)
Pabst, M. et al. Rhamnogalacturonan II structure shows variation in the side chains monosaccharide composition and methylation status within and across different plant species. Plant J. 76, 61–72 (2013)
Cartmell, A. et al. The structure and function of an arabinan-specific α-1,2-arabinofuranosidase identified from screening the activities of bacterial GH43 glycoside hydrolases. J. Biol. Chem. 286, 15483–15495 (2011)
Glenwright, A. J. et al. Structural basis for nutrient acquisition by dominant members of the human gut microbiota. Nature 541, 407–411 (2017)
Bourlard, T., Pellerin, P. & Morvan, C. Rhamnogalacturonans I and II are pectic substrates for flax-cell methyltransferases. Plant Physiol. Biochem. 35, 623–629 (1997)
Glushka, J. N. et al. Primary structure of the 2-O-methyl-α-l-fucose-containing side chain of the pectic polysaccharide, rhamnogalacturonan II. Carbohydr. Res. 338, 341–352 (2003)
Raman, R. et al. Advancing glycomics: implementation strategies at the Consortium for Functional Glycomics. Glycobiology 16, 82R–90R (2006)
Buffetto, F. et al. Recovery and fine structure variability of RGII sub-domains in wine (Vitis vinifera Merlot). Ann. Bot. 114, 1327–1337 (2014)
Chauvin, A. L., Nepogodiev, S. A. & Field, R. A. Synthesis of an apiose-containing disaccharide fragment of rhamnogalacturonan-II and some analogues. Carbohydr. Res. 339, 21–27 (2004)
Chauvin, A. L., Nepogodiev, S. A. & Field, R. A. Synthesis of a 2,3,4-triglycosylated rhamnoside fragment of rhamnogalacturonan-II side chain A using a late stage oxidation approach. J. Org. Chem. 70, 960–966 (2005)
Nepogodiev, S. A., Fais, M., Hughes, D. L. & Field, R. A. Synthesis of apiose-containing oligosaccharide fragments of the plant cell wall: fragments of rhamnogalacturonan-II side chains A and B, and apiogalacturonan. Org. Biomol. Chem. 9, 6670–6684 (2011)
Charnock, S. J. et al. The X6 “thermostabilizing” domains of xylanases are carbohydrate-binding modules: structure and biochemistry of the Clostridium thermocellum X6b domain. Biochemistry 39, 5013–5021 (2000)
Matsui, I. et al. Subsite structure of Saccharomycopsis α-amylase secreted from Saccharomyces cerevisiae. J. Biochem. 109, 566–569 (1991)
Koropatkin, N. M., Martens, E. C., Gordon, J. I. & Smith, T. J. Starch catabolism by a prominent human gut symbiont is directed by the recognition of amylose helices. Structure 16, 1105–1115 (2008)
Kabsch, W. Xds. Acta Crystallogr. D 66, 125–132 (2010)
Waterman, D. G. et al. Diffraction-geometry refinement in the DIALS framework. Acta Crystallogr. D 72, 558–575 (2016)
Winter, G. xia2: an expert system for macromolecular crystallography data reduction. J. Appl. Crystallogr. 43, 186–190 (2010)
Evans, P. R. & Murshudov, G. N. How good are my data and what is the resolution? Acta Crystallogr. D 69, 1204–1214 (2013)
Pape, T. & Schneider, T. R. HKL2MAP: a graphical user interface for macromolecular phasing with SHELX programs. J. Appl. Crystallogr. 37, 843–844 (2004)
Sheldrick, G. M. Experimental phasing with SHELXC/D/E: combining chain tracing with density modification. Acta Crystallogr. D 66, 479–485 (2010)
Cowtan, K. Fitting molecular fragments into electron density. Acta Crystallogr. D 64, 83–89 (2008)
Langer, G., Cohen, S. X., Lamzin, V. S. & Perrakis, A. Automated macromolecular model building for X-ray crystallography using ARP/wARP version 7. Nat. Protocols 3, 1171–1179 (2008)
Vagin, A. & Teplyakov, A. MOLREP: an automated program for molecular replacement. J. Appl. Crystallogr. 30, 1022–1025 (1997)
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007)
Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D 60, 2126–2132 (2004)
Vagin, A. A. et al. REFMAC5 dictionary: organization of prior chemical knowledge and guidelines for its use. Acta Crystallogr. D 60, 2184–2195 (2004)
Chen, V. B. et al. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. D 66, 12–21 (2010)
Terrapon, N., Lombard, V., Gilbert, H. J. & Henrissat, B. Automatic prediction of polysaccharide utilization loci in Bacteroidetes species. Bioinformatics 31, 647–655 (2015)
Terrapon, N., Weiner, J., Grath, S., Moore, A. D. & Bornberg-Bauer, E. Rapid similarity search of proteins using alignments of domain arrangements. Bioinformatics 30, 274–281 (2014)
Acknowledgements
This work was supported in part by a grant to H.J.G. and B.H. from the European Research Council (grant no. 322820). B.H. was also funded by Agence Nationale de la Recherche under grant number ANR 12-BIME-0006-01. H.J.G was also supported by Biotechnology and Biological Research Council (grant numbers BB/K020358/1 and BB/K001949/1), the Wellcome Trust (grant no. WT097907MA) and, with X.Z., M.-C.R. and F.B., was funded by the European Union Seventh Framework Programme under the WallTraC project (grant agreement number 263916). M.A.O and B.R.U. were supported in part by grant DE-FG02-12ER16324 from The Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the US Department of Energy. I.V. was in receipt of a Marie Skłodowska-Curie Fellowship (grant no. 707922). G.J.D. is a Royal Society Ken Murray Research Professor. D.W.A. was supported by a grant from the Beef and Cattle Research Council (FDE.15.13). We thank Diamond Light Source for access to beamline I02, I04-1 and I24 (mx1960, mx7854 and mx9948) that contributed to the results presented here, and to T. Doco and S. J. Charnock who supplied the partially purified apple RG-II.
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Contributions
Enzyme characterization was by D.N., A.R., A.C., A.S.L., I.V., A.L., D.W.A., Y.Z. and X.Z. Crystallographic studies were by A.C., A.B., A.S.L., D.N. and I.V. Purification of RG-II and oligosaccharide products was by M.A.O., A.R., D.N., A.C, A.L., A.S.L., F.B. and M.-C.R. HPLC analysis was by A.R., D.N., A.C. and A.S.L., while mass spectrometry analysis was by A.R. and J.G. Chemical synthesis was by S.N. and R.A.F. Growth analysis on purified RG-II performed by D.N. and A.R. Gene deletion strains were created and characterized by D.N and I.V. Bioinformatics and genomic annotation were by N.T. and B.H. Bacterial growth and transcriptomic experiments were by J.B., Y.X. and E.C.M. Experiments were designed by D.N., A.R., A.C., A.S.L., E.C.M. and H.J.G. The manuscript was written by H.J.G. with contributions from G.J.D., B.H., M.A.O, B.R.U., E.C.M. and W.S.Y. Figures were prepared by A.R., A.L., D.N. and A.S.L.
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Extended data figures and tables
Extended Data Figure 1 Sequence conservation and genomic organization of components of RG-II PUL1 across Bacteroidetes species.
a, PULs activated by RG-II. Genes encoding proteins of known or predicted functionalities are colour-coded. Arrows indicate the orientation of each gene. b, For Bacteroidetes species (in rows), xenologues of B. thetaiotaomicron (Bt) RG-II PUL1 proteins (in columns) were identified by reciprocal best-BLASTP hits. These organisms were coloured to reflect growth on RG-II; blue, growth in 24 h; orange, growth in 48 h; red, no growth; black, growth on RG-II was not evaluated. B. thetaiotaomicron proteins that contribute to RG-II degradation are grey. CE, carbohydrate esterase; GHXX, glycoside hydrolase family; PL1, polysaccharide lyase family. b, i, Xenologues are in the same column as the corresponding B. thetaiotaomicron protein, and percentage identity is in a red colour-scale to 24% identity. b, ii, Genomic organization of xenologues into gene clusters/loci. The top row shows 55 B. thetaiotaomicron proteins numbered according to relative position of the gene on the genome. The xenologues of the B. thetaiotaomicron genes are numbered as they appear in their respective cluster/locus. For each species, B. thetaiotaomicron RG-II PUL1 was split into separate clusters when the contiguous xenologues were separated by ≥30 unrelated genes. Each cluster has a distinct background colour (green, blue, pink and yellow). Proteins in B. xylanisolvens XB1A, not annotated previously as open-reading frames identified here by TBLASTN, are marked with a dagger. Split proteins in B. cellulosilyticus DSM 14838 due to incomplete genome assembly were numbered based on the B. cellulosilyticus WH2 genome (marked with an asterisk). B. thetaiotaomicron enzymes that were split into two distinct single module enzymes in other species are denoted with a ‘plus’ sign in the PUL. Supplementary Fig. 3 shows a complete depiction of the PUL organization in the selected species.
Extended Data Figure 2 The generation of RG-II derived oligosaccharides by mutants of B. thetaiotaomicron.
a, The site of action of the enzyme that was eliminated or not functional in the corresponding mutant. b, The structures of the oligosaccharides generated by each mutant. These molecules were isolated by size-exclusion chromatography and used as substrates to determine the mechanism of RG-II degradation. c, The structure of bespoke chemically synthesized oligosaccharides that were used as substrates to dissect the mechanism of RG-II depolymerization.
Extended Data Figure 3 Analysis of the disassembly of chains C and D.
Reactions were carried out using standard conditions as described. Control denotes that the glycan was not enzyme treated. a, Wild-type BT1013 and BT1020 were incubated with RG-II and the oligosaccharides generated by the B. thetaiotaomicron Δbt1020 mutant grown for 50 h, respectively, and the reactions were analysed by TLC. b, Wild-type and mutants of BT1013 were incubated with RG-II, and the products were subjected to HPAEC-PAD analysis. The BT1013 mutants ΔGH78 E496A and ΔGH33 Y1257A are mutants of the predicted catalytic residues of the GH78 rhamnosidase and GH33 sialidase catalytic modules, respectively. c, Δbt1020 oligosaccharide was incubated with wild-type BT1020, and the products were analysed by HPAEC-PAD and mass spectrometry. d, The activity of the N-terminal (residues Asp26 to Asp613) and C-terminal (residues Lys614 to Leu1107) regions of BT1020 were compared with the wild-type enzyme using HPAEC-PAD analysis. Arabinose and rhamnose were identified through HPAEC-PAD by co-migration with the appropriate standard, while the identity of DHA was consistent with the mass spectrometry data. The data displayed are examples from biological replicates n = 3.
Extended Data Figure 4 Crystal structures of BT1020 and BT3662.
a, b, The structures of BT1020 (a) and BT3662 (b). a, i, Schematic of BT1020, in which the domains from the N to C termini are coloured cyan (DHA hydrolase catalytic domain), blue (structural β-sandwich domain 1), yellow (β-l-arabinofuranosidase catalytic domain), salmon (structural β-sandwich domain 2). a, ii, iii, Key active site residues in stick format of the DHA hydrolase (ii) and β-l-arabinofuranosidase (iii) catalytic domains. In a, ii, the BT1020 residues (carbon and lettering coloured cyan) are overlayed with the GH33 Clostridium perfringens sialidase NanI (PDB code 2VK7; carbons and lettering are light grey and black, respectively). The ligand, N-acetyl-neuramic acid (yellow), is derived from the NanI structure. In a, iii, the active site amino acids (carbons in yellow) that interact with the bound l-Araf (carbons in green) are shown with polar contacts depicted by broken black lines. a, iv, v, Charged surface representations of the active site pockets of the DHA hydrolase (iv) and β-l-arabinofuranosidase (v) catalytic domains (with l-Araf bound to the β-l-arabinofuranosidase). Note the highly basic feature at the active sites consistent with the negatively charged DHA located in the active site or +1 subsite of the two catalytic domains. b, i, Schematic of BT3662, in which the five-bladed β-propeller catalytic domain and the β-sandwich domain are shown in green and red, respectively. In b, ii, key residues are shown in stick format. The carbons of the amino acids coloured salmon pink are catalytic residues, the yellow tyrosine is positioned in the +1 subsite, and the blue arginine residues are in the distal regions that interact with the d-GalA-containing backbone. b, iii, Solvent-exposed surface representation of b, ii, using the same colour format for the amino acids highlighted. The active site housing the catalytic residues is located in a pocket that abuts onto a shallow channel containing the arginines and tyrosine.
Extended Data Figure 5 Mechanism by which B. thetaiotaomicron depolymerizes chain B of RG-II.
The oligosaccharides generated by Δbt1003 and Δbt0986, RG-II and chemically synthesized molecules were used to determine which enzymes acted on chain B. a, Identified enzymes were added sequentially to chain B (isolated by mild acid treatment of RG-II). The reactions were carried out under standard conditions. The sugars released were identified and quantified by HPAEC-PAD. b, Examples of the use of mass spectrometry to monitor the enzymatic disassembly of chain B. Examples shown are from biological replicates, n = 3.
Extended Data Figure 6 Crystal structure of BT0986.
a, A schematic of the enzyme is displayed, revealing the (α/β)8-barrel catalytic domain (yellow) that is interrupted with three β-sandwich domains (blue), while the C-terminal domain (salmon) also folds into a β-sandwich. b, Active site of BT0986 bound to rhamnose. Catalytic amino acids are coloured magenta, other amino acids are blue, and the rhamnose is in yellow. c, Transition state mimic rhamnopyranose tetrazole bound in the active site of BT0986. Colouring as in c. The blue mesh surrounding the ligand represents the 2Fo − Fc electron density map (1.3 Å resolution) at 1.5σ. In b and c, the calcium ion in the active site is shown as a cyan sphere and its polar contacts with amino acids and ligands are indicated by black dashed lines. d, Chain B-derived heptasaccharide generated by the mutant bacterium Δbt0986 bound in the substrate-binding site of BT0986 shown as a surface representation. e, The interactions of the Δbt0986 heptasaccharide with BT0986. Amino acids that interact with the oligosaccharide are coloured blue and the sugars in both d and e are as depicted in Fig. 5. The blue mesh is the electron density of the oligosaccharide. f, Conformation of the arabinopyranose in chain B (green) and in the Δbt0986 heptasaccharide (cyan). The carbons are numbered.
Extended Data Figure 7 Crystal structure of BT1003, BT1012 and BT1002.
All amino acids are in stick format and the position of the active site in the schematics of the respect enzymes is indicated by a black box. a, Structure of the GH127 aceric acid hydrolase. a, i, Schematic of the enzyme revealing the catalytic domain (α/α)6-barrel (red) and β-sandwich domain (blue). a, ii, Overlay of the key active site residues of the aceric acid hydrolase (red) and the GH127 β-l-arabinofuranosidase HypBA1 from Bifidobacterium longum (PDB code 3WKX) (white grey). The proposed catalytic nucleophile, Cys457 in BT1003, is conserved in the two enzymes; the catalytic acid/base in HpyBA1 (Glu322), however, is a glutamine (Gln365) in the aceric acid hydrolase. The ligand (yellow) is β-l-Araf derived from the crystal structure of the β-l-arabinofuranosidase. a, iii, Structure of aceric acid (α-l-AcefA). b, Structure of the apiosidase BT1012. b, i, Schematic of the enzyme, revealing the (α/β)8-barrel catalytic domain (beige) and the C-terminal β-sandwich domain (blue). b, ii, The yellow amino acids denote the key residues in the active site (−1 subsite) that are proposed to have a direct catalytic role (Asp187 and Glu284) or substrate binding function (Gln239). The pair of arginine residues (blue) in the +1 subsite are likely to contribute to d-GalA binding through interactions with the carboxylate of the uronic acid. The aromatic residues (green) are in the −2 subsite. b, iii, Solved-exposed surface representation of the substrate-binding region of BT1012. The highlighted residues are coloured as in b, ii. c, Structure of BT1002. c, i, Schematic of the enzyme, revealing the C-terminal β-parallel helical catalytic domain (green) and the N-terminal β-sandwich domain (yellow). c, ii, Overlay of BT1002 and its closest structural homologue, a GH120 β-xylosidase (PDB code 3VSU), highlighting the position of the catalytic amino acids (green, BT1002; or light grey, β-xylosidase). The xylose in the active site of the β-xylosidase is shown to orientate the close but not identical position of the catalytic centre of the two enzymes. c, iii, Solvent-exposed surface representation of the active site pocket of BT1002, in which the catalytic residues are shown in stick format and their location on the surface depicted in red. The pocket is elongated, hinting that the 2-O-methyl-xylose appended to the l-fucose may be housed in the catalytic centre of the α-l-fucosidase.
Extended Data Figure 8 Identification of the enzymes that disassemble the backbone of RG-II.
a, BT1023 was incubated with RG-II or homogalacturonan HGA or HGB, which are 30% and 80% methyl-esterified, respectively (1% w/v), in 50 mM CAPSO buffer, pH 9.0, containing 2 mM CaCl2. RG-II substrate was either untreated or had been previously incubated with BT1013 and/or BT1020. The reactions were analysed by TLC. Control lanes denote the absence of enzyme. b, The oligosaccharide generated by the Δbt1017 mutant (Δbt1017 oligo) was incubated with BT1017 and the reaction product was analysed by mass spectrometry. c, Sequential degradation of the product generated in b by the enzymes indicated under standard conditions. The sugars released by these other enzymes (reactions 1–4) were identified and quantified by HPAEC-PAD. The structure of the oligosaccharide sugars followed the notation described in Fig. 1 and Extended Data Fig. 2. The example shown are from technical replicates n = 3.
Extended Data Figure 9 The influence of borate on the retaining apiosidase BT1012.
a, The chemical structure of apiose (top), and when the sugar is cross-linked by borate in chain A of RG-II (bottom). b, Oligosaccharide generated by the B. thetaiotaomicron mutant Δbt1010 (Δbt1010 oligo) was incubated with BT1012 under standard conditions in the presence and absence of 50 mM borate, and the products were analysed by TLC. c, The same experiment as in a except that the substrate was l-Rhap-β1,3′-d-Apif-α1,2-d-GalA-Me (Rha-Api-GalA-Me). d, Mass spectrometric analysis of the control sample and the two reactions containing BT1012 loaded onto the TLC in b. e, BT1012 was incubated under standard conditions with Rha-β1,3′-Api-α1,2-GalA-Me in the presence and absence of 2.5 M methanol. The reactions were analysed by HPAEC-PAD and mass spectrometry. The example shown are from technical replicates n = 3.
Extended Data Figure 10 Modification of the structure of RG-II.
a, Recombinant BT1001 and BT1012 were incubated with bespoke chemically synthesized oligosaccharides using standard conditions. b, The enzymatic disassembly of RG-II was used to investigate the structure of RG-II. Mass spectrometry combined with HPAEC-PAD were used to analyse the structure of the oligosaccharides generated by the mutants Δbt1010/Δbt1021 and Δbt1010/Δbt0986 before and after treatment with recombinant BT1021 using standard conditions. The enzyme reactions were analysed by HPAEC-PAD.
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Ndeh, D., Rogowski, A., Cartmell, A. et al. Complex pectin metabolism by gut bacteria reveals novel catalytic functions. Nature 544, 65–70 (2017). https://doi.org/10.1038/nature21725
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DOI: https://doi.org/10.1038/nature21725
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