1
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Santema LL, Rozeboom HJ, Borger VP, Kaya SG, Fraaije MW. Identification of a robust bacterial pyranose oxidase which displays an unusual pH dependence. J Biol Chem 2024:107885. [PMID: 39395808 DOI: 10.1016/j.jbc.2024.107885] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/16/2024] [Revised: 09/30/2024] [Accepted: 10/06/2024] [Indexed: 10/14/2024] Open
Abstract
Pyranose oxidases are valuable biocatalysts, yet only a handful of bacterial pyranose oxidases are known. These bacterial enzymes exhibit noteworthy distinctions from their extensively characterized fungal counterparts, encompassing variations in substrate specificity and structural attributes. Herein a bacterial pyranose oxidase from Oscillatoria princeps (OPOx) was biochemically characterized in detail. In contrast to the fungal pyranose oxidases, OPOx could be well expressed in Escherichia coli as soluble, fully flavinylated and active oxidase. It was found to be highly thermostable (melting temperature >90 ⁰C) and showed activity on glucose, exhibiting an exceptionally low KM value (48 μM). Elucidation of its crystal structure revealed similarities with fungal pyranose oxidases, such as being a tetramer with a large central void leading to a narrow substrate access tunnel. In the active site, the FAD cofactor is covalently bound to a histidine. OPOx displays a relatively narrow pH optimum for activity with a sharp decline at relatively basic pH values which is accompanied with a drastic change in its flavin absorbance spectrum. The pH-dependent switch in flavin absorbance features and oxidase activity was shown to be fully reversible. It is hypothesized that a glutamic acid helps to stabilize the protonated form of the histidine that is tethered to the FAD. OPOx presents itself as a valuable biocatalyst as it is highly robust, well-expressed in E. coli, shows low KM values for monosaccharides and has a peculiar pH dependent "on-off switch".
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Affiliation(s)
- Lars L Santema
- Molecular Enzymology, University of Groningen, Nijenborgh 3, 9747AG Groningen, The Netherlands
| | - Henriëtte J Rozeboom
- Molecular Enzymology, University of Groningen, Nijenborgh 3, 9747AG Groningen, The Netherlands
| | - Veronica P Borger
- Molecular Enzymology, University of Groningen, Nijenborgh 3, 9747AG Groningen, The Netherlands
| | - Saniye G Kaya
- Molecular Enzymology, University of Groningen, Nijenborgh 3, 9747AG Groningen, The Netherlands
| | - Marco W Fraaije
- Molecular Enzymology, University of Groningen, Nijenborgh 3, 9747AG Groningen, The Netherlands.
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2
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Furlanetto V, Kalyani DC, Kostelac A, Puc J, Haltrich D, Hällberg BM, Divne C. Structural and Functional Characterization of a Gene Cluster Responsible for Deglycosylation of C-glucosyl Flavonoids and Xanthonoids by Deinococcus aerius. J Mol Biol 2024; 436:168547. [PMID: 38508304 DOI: 10.1016/j.jmb.2024.168547] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/12/2024] [Revised: 03/13/2024] [Accepted: 03/14/2024] [Indexed: 03/22/2024]
Abstract
Plant C-glycosylated aromatic polyketides are important for plant and animal health. These are specialized metabolites that perform functions both within the plant, and in interaction with soil or intestinal microbes. Despite the importance of these plant compounds, there is still limited knowledge of how they are metabolized. The Gram-positive aerobic soil bacterium Deinococcus aerius strain TR0125 and other Deinococcus species thrive in a wide range of harsh environments. In this work, we identified a C-glycoside deglycosylation gene cluster in the genome of D. aerius. The cluster includes three genes coding for a GMC-type oxidoreductase (DaCGO1) that oxidizes the glucosyl C3 position in aromatic C-glucosyl compounds, which in turn provides the substrate for the C-glycoside deglycosidase (DaCGD; composed of α+β subunits) that cleaves the glucosyl-aglycone C-C bond. Our results from size-exclusion chromatography, single particle cryo-electron microscopy and X-ray crystallography show that DaCGD is an α2β2 heterotetramer, which represents a novel oligomeric state among bacterial CGDs. Importantly, the high-resolution X-ray structure of DaCGD provides valuable insights into the activation of the catalytic hydroxide ion by Lys261. DaCGO1 is specific for the 6-C-glucosyl flavones isovitexin, isoorientin and the 2-C-glucosyl xanthonoid mangiferin, and the subsequent C-C-bond cleavage by DaCGD generated apigenin, luteolin and norathyriol, respectively. Of the substrates tested, isovitexin was the preferred substrate (DaCGO1, Km 0.047 mM, kcat 51 min-1; DaCGO1/DaCGD, Km 0.083 mM, kcat 0.42 min-1).
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Affiliation(s)
- Valentina Furlanetto
- School of Engineering Sciences in Chemistry, Biotechnology, and Health, CBH, KTH Royal Institute of Technology, 100 44 Stockholm, Sweden
| | - Dayanand C Kalyani
- School of Engineering Sciences in Chemistry, Biotechnology, and Health, CBH, KTH Royal Institute of Technology, 100 44 Stockholm, Sweden
| | - Anja Kostelac
- Laboratory of Food Biotechnology, Department of Food Science and Technology, BOKU-University of Natural Resources and Life Sciences, 1190 Vienna, Austria; Doctoral Programme BioToP-Biomolecular Technology of Proteins, BOKU-University of Natural Resources and Life Sciences, 1180 Vienna, Austria
| | - Jolanta Puc
- Laboratory of Food Biotechnology, Department of Food Science and Technology, BOKU-University of Natural Resources and Life Sciences, 1190 Vienna, Austria
| | - Dietmar Haltrich
- Laboratory of Food Biotechnology, Department of Food Science and Technology, BOKU-University of Natural Resources and Life Sciences, 1190 Vienna, Austria
| | - B Martin Hällberg
- Department of Cell and Molecular Biology, Karolinska Institutet, 171 77 Stockholm, Sweden
| | - Christina Divne
- School of Engineering Sciences in Chemistry, Biotechnology, and Health, CBH, KTH Royal Institute of Technology, 100 44 Stockholm, Sweden.
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3
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Taborda A, Frazão T, Rodrigues MV, Fernández-Luengo X, Sancho F, Lucas MF, Frazão C, Melo EP, Ventura MR, Masgrau L, Borges PT, Martins LO. Mechanistic insights into glycoside 3-oxidases involved in C-glycoside metabolism in soil microorganisms. Nat Commun 2023; 14:7289. [PMID: 37963862 PMCID: PMC10646112 DOI: 10.1038/s41467-023-42000-3] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/13/2023] [Accepted: 09/27/2023] [Indexed: 11/16/2023] Open
Abstract
C-glycosides are natural products with important biological activities but are recalcitrant to degradation. Glycoside 3-oxidases (G3Oxs) are recently identified bacterial flavo-oxidases from the glucose-methanol-coline (GMC) superfamily that catalyze the oxidation of C-glycosides with the concomitant reduction of O2 to H2O2. This oxidation is followed by C-C acid/base-assisted bond cleavage in two-step C-deglycosylation pathways. Soil and gut microorganisms have different oxidative enzymes, but the details of their catalytic mechanisms are largely unknown. Here, we report that PsG3Ox oxidizes at 50,000-fold higher specificity (kcat/Km) the glucose moiety of mangiferin to 3-keto-mangiferin than free D-glucose to 2-keto-glucose. Analysis of PsG3Ox X-ray crystal structures and PsG3Ox in complex with glucose and mangiferin, combined with mutagenesis and molecular dynamics simulations, reveal distinctive features in the topology surrounding the active site that favor catalytically competent conformational states suitable for recognition, stabilization, and oxidation of the glucose moiety of mangiferin. Furthermore, their distinction to pyranose 2-oxidases (P2Oxs) involved in wood decay and recycling is discussed from an evolutionary, structural, and functional viewpoint.
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Grants
- EC | Horizon 2020 Framework Programme (EU Framework Programme for Research and Innovation H2020)
- Fundação para a Ciência e Tecnologia, Portugal, grants 2022.02027.PTDC, UIDB/04612/2020 and UIDP/04612/2020, LA/P/0087/2020, PTDC/BII-BBF/29564/2017, and AAC 01/SAICT/2016 Fundação para a Ciência e Tecnologia, Portugal, Ph.D. fellowships 2020.07928, 2022.13872, and 2022.09426 Ministry of Science and Innovation, Spain, grant PID2021-126897NB-I00 and fellowship PRE2019-088412, funded by the MCIN/AEI/10.13039/501100011033/ FEDER, EU
- Fundação para a Ciência e Tecnologia (FCT), Portugal, grants UIDB/04326/2020, UIDP/043226/2020 and LA/P/0101/2020
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Affiliation(s)
- André Taborda
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal
| | - Tomás Frazão
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal
| | - Miguel V Rodrigues
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal
| | | | - Ferran Sancho
- Zymvol Biomodeling, C/ Pau Claris, 94, 3B, 08010, Barcelona, Spain
| | | | - Carlos Frazão
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal
| | - Eduardo P Melo
- Centro de Ciências do Mar, Universidade do Algarve, 8005-139, Faro, Portugal
| | - M Rita Ventura
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal
| | - Laura Masgrau
- Department of Chemistry, Universitat Autònoma de Barcelona, 08193, Bellaterra, Spain
- Zymvol Biomodeling, C/ Pau Claris, 94, 3B, 08010, Barcelona, Spain
| | - Patrícia T Borges
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal
| | - Lígia O Martins
- Instituto de Tecnologia Química e Biológica António Xavier, Universidade Nova de Lisboa, Av da República, 2780-157, Oeiras, Portugal.
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4
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Savino S, Fraaije MW. The vast repertoire of carbohydrate oxidases: An overview. Biotechnol Adv 2020; 51:107634. [PMID: 32961251 DOI: 10.1016/j.biotechadv.2020.107634] [Citation(s) in RCA: 10] [Impact Index Per Article: 2.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/03/2020] [Revised: 08/12/2020] [Accepted: 09/06/2020] [Indexed: 01/01/2023]
Abstract
Carbohydrates are widely abundant molecules present in a variety of forms. For their biosynthesis and modification, nature has evolved a plethora of carbohydrate-acting enzymes. Many of these enzymes are of particular interest for biotechnological applications, where they can be used as biocatalysts or biosensors. Among the enzymes catalysing conversions of carbohydrates are the carbohydrate oxidases. These oxidative enzymes belong to different structural families and use different cofactors to perform the oxidation reaction of CH-OH bonds in carbohydrates. The variety of carbohydrate oxidases available in nature reflects their specificity towards different sugars and selectivity of the oxidation site. Thanks to their properties, carbohydrate oxidases have received a lot of attention in basic and applied research, such that nowadays their role in biotechnological processes is of paramount importance. In this review we provide an overview of the available knowledge concerning the known carbohydrate oxidases. The oxidases are first classified according to their structural features. After a description on their mechanism of action, substrate acceptance and characterisation, we report on the engineering of the different carbohydrate oxidases to enhance their employment in biocatalysis and biotechnology. In the last part of the review we highlight some practical applications for which such enzymes have been exploited.
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Affiliation(s)
- Simone Savino
- Molecular Enzymology Group, University of Groningen, Nijenborgh 4, 9747AG Groningen, the Netherlands
| | - Marco W Fraaije
- Molecular Enzymology Group, University of Groningen, Nijenborgh 4, 9747AG Groningen, the Netherlands.
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5
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Sriwaiyaphram K, Punthong P, Sucharitakul J, Wongnate T. Structure and function relationships of sugar oxidases and their potential use in biocatalysis. Enzymes 2020; 47:193-230. [PMID: 32951824 DOI: 10.1016/bs.enz.2020.05.006] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/24/2023]
Abstract
Several sugar oxidases that catalyze the oxidation of sugars have been isolated and characterized. These enzymes can be classified as flavoenzyme due to the presence of flavin adenine dinucleotide (FAD) as a cofactor. Sugar oxidases have been proposed to be the key biocatalyst in biotransformation of carbohydrates which can potentially convert sugars to provide a pool of intermediates for synthesis of rare sugars, fine chemicals and drugs. Moreover, sugar oxidases have been applied in biosensing of various biomolecules in food industries, diagnosis of diseases and environmental pollutant detection. This review provides the discussions on general properties, current mechanistic understanding, structural determination, biocatalytic application, and biosensor integration of representative sugar oxidase enzymes, namely pyranose 2-oxidase (P2O), glucose oxidase (GO), hexose oxidase (HO), and oligosaccharide oxidase. The information regarding the relationship between structure and function of these sugar oxidases points out the key properties of this particular group of enzymes that can be modified by engineering, which had resulted in a remarkable economic importance.
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Affiliation(s)
- Kanokkan Sriwaiyaphram
- School of Biomolecular Science and Engineering, Vidyasirimedhi Institute of Science and Technology (VISTEC), Rayong, Thailand
| | - Pangrum Punthong
- School of Biomolecular Science and Engineering, Vidyasirimedhi Institute of Science and Technology (VISTEC), Rayong, Thailand
| | - Jeerus Sucharitakul
- Department of Biochemistry, Faculty of Dentistry, Chulalongkorn University, Bangkok, Thailand
| | - Thanyaporn Wongnate
- School of Biomolecular Science and Engineering, Vidyasirimedhi Institute of Science and Technology (VISTEC), Rayong, Thailand.
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6
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Wongnate T, Surawatanawong P, Chuaboon L, Lawan N, Chaiyen P. The Mechanism of Sugar C−H Bond Oxidation by a Flavoprotein Oxidase Occurs by a Hydride Transfer Before Proton Abstraction. Chemistry 2019; 25:4460-4471. [DOI: 10.1002/chem.201806078] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/06/2018] [Revised: 01/16/2019] [Indexed: 11/06/2022]
Affiliation(s)
- Thanyaporn Wongnate
- School of Biomolecular Science & EngineeringVidyasirimedhi Institute of Science and Technology (VISTEC), Wangchan Valley Rayong 21210 Thailand
| | - Panida Surawatanawong
- Department of Chemistry and Center of Excellence, for Innovation in ChemistryMahidol University Bangkok 10400 Thailand
| | - Litavadee Chuaboon
- Department of Biochemistry and Center for Excellence, in Protein and Enzyme Technology, Faculty of ScienceMahidol University Bangkok 10400 Thailand
| | - Narin Lawan
- Department of Chemistry, Faculty of ScienceChiang Mai University Chiang Mai 50200 Thailand
| | - Pimchai Chaiyen
- School of Biomolecular Science & EngineeringVidyasirimedhi Institute of Science and Technology (VISTEC), Wangchan Valley Rayong 21210 Thailand
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7
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Analysis of Agaricus meleagris pyranose dehydrogenase N-glycosylation sites and performance of partially non-glycosylated enzymes. Enzyme Microb Technol 2017; 99:57-66. [PMID: 28193332 DOI: 10.1016/j.enzmictec.2017.01.008] [Citation(s) in RCA: 5] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/30/2016] [Revised: 01/17/2017] [Accepted: 01/20/2017] [Indexed: 11/22/2022]
Abstract
Pyranose Dehydrogenase 1 from the basidiomycete Agaricus meleagris (AmPDH1) is an oxidoreductase capable of oxidizing a broad variety of sugars. Due to this and its ability of dioxidation of substrates and no side production of hydrogen peroxide, it is studied for use in enzymatic bio-fuel cells. In-vitro deglycosylated AmPDH1 as well as knock-out mutants of the N-glycosylation sites N75 and N175, near the active site entrance, were previously shown to improve achievable current densities of graphite electrodes modified with AmPDH1 and an osmium redox polymer acting as a redox mediator, up to 10-fold. For a better understanding of the role of N-glycosylation of AmPDH1, a systematic set of N-glycosylation site mutants was investigated in this work, regarding expression efficiency, enzyme activity and stability. Furthermore, the site specific extend of N-glycosylation was compared between native and recombinant wild type AmPDH1. Knocking out the site N252 prevented the attachment of significantly extended N-glycan structures as detected on polyacrylamide gel electrophoresis, but did not significantly alter enzyme performance on modified electrodes. This suggests that not the molecule size but other factors like accessibility of the active site improved performance of deglycosylated AmPDH1/osmium redox polymer modified electrodes. A fourth N-glycosylation site of AmPDH1 could be confirmed by mass spectrometry at N319, which appeared to be conserved in related fungal pyranose dehydrogenases but not in other members of the glucose-methanol-choline oxidoreductase structural family. This site was shown to be the only one that is essential for functional recombinant expression of the enzyme.
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8
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Graf MMH, Sucharitakul J, Bren U, Chu DB, Koellensperger G, Hann S, Furtmüller PG, Obinger C, Peterbauer CK, Oostenbrink C, Chaiyen P, Haltrich D. Reaction of pyranose dehydrogenase from Agaricus meleagris with its carbohydrate substrates. FEBS J 2015; 282:4218-41. [PMID: 26284701 PMCID: PMC4950071 DOI: 10.1111/febs.13417] [Citation(s) in RCA: 13] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/22/2015] [Revised: 08/04/2015] [Accepted: 08/13/2015] [Indexed: 01/25/2023]
Abstract
Monomeric Agaricus meleagris pyranose dehydrogenase (AmPDH) belongs to the glucose-methanol-choline family of oxidoreductases. An FAD cofactor is covalently tethered to His103 of the enzyme. AmPDH can double oxidize various mono- and oligosaccharides at different positions (C1 to C4). To study the structure/function relationship of selected active-site residues of AmPDH pertaining to substrate (carbohydrate) turnover in more detail, several active-site variants were generated, heterologously expressed in Pichia pastoris, and characterized by biochemical, biophysical and computational means. The crystal structure of AmPDH shows two active-site histidines, both of which could take on the role as the catalytic base in the reductive half-reaction. Steady-state kinetics revealed that His512 is the only catalytic base because H512A showed a reduction in (kcat /KM )glucose by a factor of 10(5) , whereas this catalytic efficiency was reduced by two or three orders of magnitude for His556 variants (H556A, H556N). This was further corroborated by transient-state kinetics, where a comparable decrease in the reductive rate constant was observed for H556A, whereas the rate constant for the oxidative half-reaction (using benzoquinone as substrate) was increased for H556A compared to recombinant wild-type AmPDH. Steady-state kinetics furthermore indicated that Gln392, Tyr510, Val511 and His556 are important for the catalytic efficiency of PDH. Molecular dynamics (MD) simulations and free energy calculations were used to predict d-glucose oxidation sites, which were validated by GC-MS measurements. These simulations also suggest that van der Waals interactions are the main driving force for substrate recognition and binding.
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Affiliation(s)
- Michael M H Graf
- Food Biotechnology Laboratory, Department of Food Science and Technology, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
| | - Jeerus Sucharitakul
- Department of Biochemistry, Faculty of Dentistry, Chulalongkorn University, Bangkok, Thailand
| | - Urban Bren
- Institute of Molecular Modeling and Simulation, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
- Laboratory for Physical Chemistry and Chemical Thermodynamics, Faculty of Chemistry and Chemical Technology, University of Maribor, Slovenia
| | - Dinh Binh Chu
- Division of Analytical Chemistry, Department of Chemistry, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
- School of Chemical Engineering, Department of Analytical Chemistry, Hanoi University of Science and Technology, Hanoi, Vietnam
| | - Gunda Koellensperger
- Institute of Analytical Chemistry, Faculty of Chemistry, University of Vienna, Austria
| | - Stephan Hann
- Division of Analytical Chemistry, Department of Chemistry, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
| | - Paul G Furtmüller
- Division of Biochemistry, Department of Chemistry, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
| | - Christian Obinger
- Division of Biochemistry, Department of Chemistry, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
| | - Clemens K Peterbauer
- Food Biotechnology Laboratory, Department of Food Science and Technology, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
| | - Chris Oostenbrink
- Institute of Molecular Modeling and Simulation, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
| | - Pimchai Chaiyen
- Department of Biochemistry and Center of Excellence in Protein Structure and Function, Faculty of Science, Mahidol University, Bangkok, Thailand
| | - Dietmar Haltrich
- Food Biotechnology Laboratory, Department of Food Science and Technology, University of Natural Resources and Life Sciences Vienna (BOKU), Austria
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9
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Pyranose dehydrogenase ligand promiscuity: a generalized approach to simulate monosaccharide solvation, binding, and product formation. PLoS Comput Biol 2014; 10:e1003995. [PMID: 25500811 PMCID: PMC4263366 DOI: 10.1371/journal.pcbi.1003995] [Citation(s) in RCA: 8] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/26/2014] [Accepted: 10/13/2014] [Indexed: 11/19/2022] Open
Abstract
The flavoenzyme pyranose dehydrogenase (PDH) from the litter decomposing fungus Agaricus meleagris oxidizes many different carbohydrates occurring during lignin degradation. This promiscuous substrate specificity makes PDH a promising catalyst for bioelectrochemical applications. A generalized approach to simulate all 32 possible aldohexopyranoses in the course of one or a few molecular dynamics (MD) simulations is reported. Free energy calculations according to the one-step perturbation (OSP) method revealed the solvation free energies (ΔGsolv) of all 32 aldohexopyranoses in water, which have not yet been reported in the literature. The free energy difference between β- and α-anomers (ΔGβ-α) of all d-stereoisomers in water were compared to experimental values with a good agreement. Moreover, the free-energy differences (ΔG) of the 32 stereoisomers bound to PDH in two different poses were calculated from MD simulations. The relative binding free energies (ΔΔGbind) were calculated and, where available, compared to experimental values, approximated from Km values. The agreement was very good for one of the poses, in which the sugars are positioned in the active site for oxidation at C1 or C2. Distance analysis between hydrogens of the monosaccharide and the reactive N5-atom of the flavin adenine dinucleotide (FAD) revealed that oxidation is possible at HC1 or HC2 for pose A, and at HC3 or HC4 for pose B. Experimentally detected oxidation products could be rationalized for the majority of monosaccharides by combining ΔΔGbind and a reweighted distance analysis. Furthermore, several oxidation products were predicted for sugars that have not yet been tested experimentally, directing further analyses. This study rationalizes the relationship between binding free energies and substrate promiscuity in PDH, providing novel insights for its applicability in bioelectrochemistry. The results suggest that a similar approach could be applied to study promiscuity of other enzymes. Generally, enzymes are perceived as being specific for both their substrates and the reaction they catalyze. This standard paradigm started to shift and currently enzyme promiscuity towards various substrates is perceived rather as the rule than the exception. Enzyme promiscuity seems to be vital for proteins to acquire new functions, and therefore for evolution itself. The driving forces for promiscuity are manifold and consequently challenging to study. Binding free energies, which can be calculated from computer simulations, represent a convenient measure for them. Here, we investigate the binding free energies between the enzyme pyranose dehydrogenase (PDH) and a sugar-substrate by computational means. PDH has an extraordinarily promiscuous substrate-specificity, making it interesting for e.g. bioelectrochemical applications. By introducing modifications to the sugar-structure used for the molecular dynamics simulations, we could simultaneously study all 32 possible aldohexopyranoses from a single simulation. This saves costly computational resources and time for setting up and analyzing the simulations. We could nicely reproduce experimental results and predict so far undetected sugar-oxidation products, directing further experiments. This study gives novel insights into PDH's substrate promiscuity and the enzyme's applicability. A similar approach could be applied to study the promiscuity of other enzymes.
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10
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Tan TC, Spadiut O, Gandini R, Haltrich D, Divne C. Structural basis for binding of fluorinated glucose and galactose to Trametes multicolor pyranose 2-oxidase variants with improved galactose conversion. PLoS One 2014; 9:e86736. [PMID: 24466218 PMCID: PMC3897772 DOI: 10.1371/journal.pone.0086736] [Citation(s) in RCA: 8] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/23/2013] [Accepted: 12/15/2013] [Indexed: 11/18/2022] Open
Abstract
Each year, about six million tons of lactose are generated from liquid whey as industrial byproduct, and optimally this large carbohydrate waste should be used for the production of value-added products. Trametes multicolor pyranose 2-oxidase (TmP2O) catalyzes the oxidation of various monosaccharides to the corresponding 2-keto sugars. Thus, a potential use of TmP2O is to convert the products from lactose hydrolysis, D-glucose and D-galactose, to more valuable products such as tagatose. Oxidation of glucose is however strongly favored over galactose, and oxidation of both substrates at more equal rates is desirable. Characterization of TmP2O variants (H450G, V546C, H450G/V546C) with improved D-galactose conversion has been given earlier, of which H450G displayed the best relative conversion between the substrates. To rationalize the changes in conversion rates, we have analyzed high-resolution crystal structures of the aforementioned mutants with bound 2- and 3-fluorinated glucose and galactose. Binding of glucose and galactose in the productive 2-oxidation binding mode is nearly identical in all mutants, suggesting that this binding mode is essentially unaffected by the mutations. For the competing glucose binding mode, enzyme variants carrying the H450G replacement stabilize glucose as the α-anomer in position for 3-oxidation. The backbone relaxation at position 450 allows the substrate-binding loop to fold tightly around the ligand. V546C however stabilize glucose as the β-anomer using an open loop conformation. Improved binding of galactose is enabled by subtle relaxation effects at key active-site backbone positions. The competing binding mode for galactose 2-oxidation by V546C stabilizes the β-anomer for oxidation at C1, whereas H450G variants stabilize the 3-oxidation binding mode of the galactose α-anomer. The present study provides a detailed description of binding modes that rationalize changes in the relative conversion rates of D-glucose and D-galactose and can be used to refine future enzyme designs for more efficient use of lactose-hydrolysis byproducts.
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Affiliation(s)
- Tien Chye Tan
- Department of Medical Biochemistry and Biophysics, Karolinska Institute, Stockholm, Sweden
| | - Oliver Spadiut
- School of Biotechnology, Royal Institute of Technology, Stockholm, Sweden
| | - Rosaria Gandini
- Department of Medical Biochemistry and Biophysics, Karolinska Institute, Stockholm, Sweden
| | - Dietmar Haltrich
- Food Biotechnology Laboratory, BOKU University of Natural Resources and Life Sciences, Vienna, Austria
| | - Christina Divne
- Department of Medical Biochemistry and Biophysics, Karolinska Institute, Stockholm, Sweden ; School of Biotechnology, Royal Institute of Technology, Stockholm, Sweden
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11
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Wongnate T, Surawatanawong P, Visitsatthawong S, Sucharitakul J, Scrutton NS, Chaiyen P. Proton-Coupled Electron Transfer and Adduct Configuration Are Important for C4a-Hydroperoxyflavin Formation and Stabilization in a Flavoenzyme. J Am Chem Soc 2013; 136:241-53. [DOI: 10.1021/ja4088055] [Citation(s) in RCA: 52] [Impact Index Per Article: 4.7] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
Affiliation(s)
- Thanyaporn Wongnate
- Department
of Biochemistry and Center of Excellence in Protein Structure and
Function, Faculty of Science, Mahidol University, Bangkok, 10400 Thailand
| | - Panida Surawatanawong
- Department
of Chemistry and Center of Excellence for Innovation in Chemistry, Mahidol University, Bangkok 10400 Thailand
| | - Surawit Visitsatthawong
- Department
of Chemistry and Center of Excellence for Innovation in Chemistry, Mahidol University, Bangkok 10400 Thailand
| | - Jeerus Sucharitakul
- Department
of Biochemistry, Faculty of Dentistry, Chulalongkorn University, Henri-Dunant
Road, Patumwan, Bangkok, 10300 Thailand
| | - Nigel S. Scrutton
- Manchester
Institute of Biotechnology and Faculty of Life Sciences, The University of Manchester, Manchester M1 7DN United Kingdom
| | - Pimchai Chaiyen
- Department
of Biochemistry and Center of Excellence in Protein Structure and
Function, Faculty of Science, Mahidol University, Bangkok, 10400 Thailand
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12
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Hassan N, Tan TC, Spadiut O, Pisanelli I, Fusco L, Haltrich D, Peterbauer CK, Divne C. Crystal structures of Phanerochaete chrysosporium pyranose 2-oxidase suggest that the N-terminus acts as a propeptide that assists in homotetramer assembly. FEBS Open Bio 2013; 3:496-504. [PMID: 24282677 PMCID: PMC3839853 DOI: 10.1016/j.fob.2013.10.010] [Citation(s) in RCA: 16] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/28/2013] [Revised: 10/30/2013] [Accepted: 10/31/2013] [Indexed: 11/17/2022] Open
Abstract
The flavin-dependent homotetrameric enzyme pyranose 2-oxidase (P2O) is found mostly, but not exclusively, in lignocellulose-degrading fungi where it catalyzes the oxidation of β-d-glucose to the corresponding 2-keto sugar concomitantly with hydrogen peroxide formation during lignin solubilization. Here, we present crystal structures of P2O from the efficient lignocellulolytic basidiomycete Phanerochaete chrysosporium. Structures were determined of wild-type PcP2O from the natural fungal source, and two variants of recombinant full-length PcP2O, both in complex with the slow substrate 3-deoxy-3-fluoro-β-d-glucose. The active sites in PcP2O and P2O from Trametes multicolor (TmP2O) are highly conserved with identical substrate binding. Our structural analysis suggests that the 17 °C higher melting temperature of PcP2O compared to TmP2O is due to an increased number of intersubunit salt bridges. The structure of recombinant PcP2O expressed with its natural N-terminal sequence, including a proposed propeptide segment, reveals that the first five residues of the propeptide intercalate at the interface between A and B subunits to form stabilizing, mainly hydrophobic, interactions. In the structure of mature PcP2O purified from the natural source, the propeptide segment in subunit A has been replaced by a nearby loop in the B subunit. We propose that the propeptide in subunit A stabilizes the A/B interface of essential dimers in the homotetramer and that, upon maturation, it is replaced by the loop in the B subunit to form the mature subunit interface. This would imply that the propeptide segment of PcP2O acts as an intramolecular chaperone for oligomerization at the A/B interface of the essential dimer. Structures of pyranose 2-oxidase from Phanerochaete chrysosporium were determined. The N-terminus may act as a propeptide with a role in homotetramer assembly. A large number of salt bridges between subunits provides thermostability. The substrate is bound in the productive binding mode for oxidation at C2.
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Key Words
- 2FGlc, 2-deoxy-2-fluoro-d-glucose
- 3FGlc, 3-deoxy-3-fluoro-d-glucose
- Crystal structure
- DTT, dithiothreitol
- HEPES, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
- IMAC, by immobilized metal ion affinity chromatography
- IPTG, β-d-1-thiogalactopyranoside
- Lignin degradation
- MES, 2-(N-morpholino) ethanesulfonic acid
- MWCO, molecular weight cut off
- Oligomerization
- P2O, pyranose oxidase
- PBS, phosphate buffered saline
- PDB, Protein Data Bank
- PEG, polyethylene glycol
- Propeptide
- Pyranose 2-oxidase
- TEV, Tobacco Etch Virus
- Thermostability
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Affiliation(s)
- Noor Hassan
- KTH Royal Institute of Technology, School of Biotechnology, Albanova University Center, Roslagstullsbacken 21, S-10691 StockholmSweden
| | - Tien-Chye Tan
- Karolinska Institute, Department of Medical Biochemistry and Biophysics, Scheelelaboratoriet, Scheeles väg 2, S-17177 StockholmSweden
| | - Oliver Spadiut
- KTH Royal Institute of Technology, School of Biotechnology, Albanova University Center, Roslagstullsbacken 21, S-10691 StockholmSweden
| | - Ines Pisanelli
- BOKU University of Natural Resources and Life Sciences, Food Biotechnology Laboratory, A-1190 ViennaAustria
| | - Laura Fusco
- BOKU University of Natural Resources and Life Sciences, Food Biotechnology Laboratory, A-1190 ViennaAustria
| | - Dietmar Haltrich
- BOKU University of Natural Resources and Life Sciences, Food Biotechnology Laboratory, A-1190 ViennaAustria
| | - Clemens K. Peterbauer
- BOKU University of Natural Resources and Life Sciences, Food Biotechnology Laboratory, A-1190 ViennaAustria
| | - Christina Divne
- KTH Royal Institute of Technology, School of Biotechnology, Albanova University Center, Roslagstullsbacken 21, S-10691 StockholmSweden
- Karolinska Institute, Department of Medical Biochemistry and Biophysics, Scheelelaboratoriet, Scheeles väg 2, S-17177 StockholmSweden
- Corresponding author at: KTH Royal Institute of Technology, School of Biotechnology, Albanova University Center, Roslagstullsbacken 21, S-10691StockholmSweden. Tel.: +46 8 55378296; fax: +46 8 55378468.
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13
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Wongnate T, Chaiyen P. The substrate oxidation mechanism of pyranose 2-oxidase and other related enzymes in the glucose-methanol-choline superfamily. FEBS J 2013; 280:3009-27. [DOI: 10.1111/febs.12280] [Citation(s) in RCA: 70] [Impact Index Per Article: 6.4] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/19/2013] [Revised: 04/01/2013] [Accepted: 04/04/2013] [Indexed: 01/13/2023]
Affiliation(s)
- Thanyaporn Wongnate
- Department of Biochemistry and Center of Excellence in Protein Structure and Function, Faculty of Science; Mahidol University; Bangkok; Thailand
| | - Pimchai Chaiyen
- Department of Biochemistry and Center of Excellence in Protein Structure and Function, Faculty of Science; Mahidol University; Bangkok; Thailand
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14
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Molecular dynamics simulations give insight into D-glucose dioxidation at C2 and C3 by Agaricus meleagris pyranose dehydrogenase. J Comput Aided Mol Des 2013; 27:295-304. [PMID: 23591812 PMCID: PMC3657087 DOI: 10.1007/s10822-013-9645-7] [Citation(s) in RCA: 26] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/13/2012] [Accepted: 04/04/2013] [Indexed: 11/11/2022]
Abstract
The flavin-dependent sugar oxidoreductase pyranose dehydrogenase (PDH) from the plant litter-degrading fungus Agaricus meleagris oxidizes d-glucose (GLC) efficiently at positions C2 and C3. The closely related pyranose 2-oxidase (P2O) from Trametes multicolor oxidizes GLC only at position C2. Consequently, the electron output per molecule GLC is twofold for PDH compared to P2O making it a promising catalyst for bioelectrochemistry or for introducing novel carbonyl functionalities into sugars. The aim of this study was to rationalize the mechanism of GLC dioxidation employing molecular dynamics simulations of GLC–PDH interactions. Shape complementarity through nonpolar van der Waals interactions was identified as the main driving force for GLC binding. Together with a very diverse hydrogen-bonding pattern, this has the potential to explain the experimentally observed promiscuity of PDH towards different sugars. Based on geometrical analysis, we propose a similar reaction mechanism as in P2O involving a general base proton abstraction, stabilization of the transition state, an alkoxide intermediate, through interaction with a protonated catalytic histidine followed by a hydride transfer to the flavin N5 atom. Our data suggest that the presence of the two potential catalytic bases His-512 and His-556 increases the versatility of the enzyme, by employing the most suitably oriented base depending on the substrate and its orientation in the active site. Our findings corroborate and rationalize the experimentally observed dioxidation of GLC by PDH and its promiscuity towards different sugars.
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15
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Mugo AN, Kobayashi J, Yamasaki T, Mikami B, Ohnishi K, Yoshikane Y, Yagi T. Crystal structure of pyridoxine 4-oxidase from Mesorhizobium loti. BIOCHIMICA ET BIOPHYSICA ACTA-PROTEINS AND PROTEOMICS 2013; 1834:953-63. [PMID: 23501672 DOI: 10.1016/j.bbapap.2013.03.004] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 12/20/2012] [Revised: 02/21/2013] [Accepted: 03/04/2013] [Indexed: 10/27/2022]
Abstract
Pyridoxine 4-oxidase (PNOX) from Mesorhizobium loti is a monomeric glucose-methanol-choline (GMC) oxidoreductase family enzyme, catalyzes FAD-dependent oxidation of pyridoxine (PN) into pyridoxal, and is the first enzyme in pathway I for the degradation of PN. The tertiary structures of PNOX with a C-terminal His6-tag and PNOX-pyridoxamine (PM) complex were determined at 2.2Å and at 2.1Å resolutions, respectively. The overall structure consisted of FAD-binding and substrate-binding domains. In the active site, His460, His462, and Pro504 were located on the re-face of the isoalloxazine ring of FAD. PM binds to the active site through several hydrogen bonds. The side chains of His462 and His460 are located at 2.7 and 3.1Å from the N4' atom of PM. The activities of His460Ala and His462Ala mutant PNOXs were very low, and 460Ala/His462Ala double mutant PNOX exhibited no activity. His462 may act as a general base for the abstraction of a proton from the 4'-hydroxyl of PN. His460 may play a role in the binding and positioning of PN. The C4' atom in PM is located at 3.2Å, and the hydride ion from the C4' atom may be transferred to the N5 atom of the isoalloxazine ring. The comparison of active site residues in GMC oxidoreductase shows that Pro504 in PNOX corresponds to Asn or His of the conserved His-Asn or His-His pair in other GMC oxidoreductases. The function of the novel proline residue was discussed.
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Affiliation(s)
- Andrew Njagi Mugo
- Graduate School of Integral Arts and Science, Kochi University, Nankoku, Kochi, Japan
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16
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Prongjit M, Sucharitakul J, Palfey BA, Chaiyen P. Oxidation mode of pyranose 2-oxidase is controlled by pH. Biochemistry 2013; 52:1437-45. [PMID: 23356577 DOI: 10.1021/bi301442x] [Citation(s) in RCA: 20] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/29/2022]
Abstract
Pyranose 2-oxidase (P2O) from Trametes multicolor is a flavoenzyme that catalyzes the oxidation of d-glucose and other aldopyranose sugars at the C2 position by using O₂ as an electron acceptor to form the corresponding 2-keto-sugars and H₂O₂. In this study, the effects of pH on the oxidative half-reaction of P2O were investigated using stopped-flow spectrophotometry. The results showed that flavin oxidation occurred via different pathways depending on the pH of the environment. At pH values lower than 8.0, reduced P2O reacts with O₂ to form a C4a-hydroperoxyflavin intermediate, leading to elimination of H₂O₂. At pH 8.0 and higher, the majority of the reduced P2O reacts with O₂ via a pathway that does not allow detection of the C4a-hydroperoxyflavin, and flavin oxidation occurs with decreased rate constants upon the rise in pH. The switching between the two modes of P2O oxidation is controlled by protonation of a group which has a pK(a) of 7.6 ± 0.1. Oxidation reactions of reduced P2O under rapid pH change as performed by stopped-flow mixing were different from the same reactions performed with enzyme pre-equilibrated at the same specified pH values, implying that the protonation of the group which controls the mode of flavin oxidation cannot be rapidly equilibrated with outside solvent. Using a double-mixing stopped-flow experiment, a rate constant for proton dissociation from the reaction site was determined to be 21.0 ± 0.4 s⁻¹.
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Affiliation(s)
- Methinee Prongjit
- Department of Biochemistry and Center of Excellence in Protein Structure and Function, Faculty of Science, Mahidol University, Bangkok 10400, Thailand
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17
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Tan TC, Spadiut O, Wongnate T, Sucharitakul J, Krondorfer I, Sygmund C, Haltrich D, Chaiyen P, Peterbauer CK, Divne C. The 1.6 Å crystal structure of pyranose dehydrogenase from Agaricus meleagris rationalizes substrate specificity and reveals a flavin intermediate. PLoS One 2013; 8:e53567. [PMID: 23326459 PMCID: PMC3541233 DOI: 10.1371/journal.pone.0053567] [Citation(s) in RCA: 42] [Impact Index Per Article: 3.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/16/2012] [Accepted: 11/29/2012] [Indexed: 11/18/2022] Open
Abstract
Pyranose dehydrogenases (PDHs) are extracellular flavin-dependent oxidoreductases secreted by litter-decomposing fungi with a role in natural recycling of plant matter. All major monosaccharides in lignocellulose are oxidized by PDH at comparable yields and efficiencies. Oxidation takes place as single-oxidation or sequential double-oxidation reactions of the carbohydrates, resulting in sugar derivatives oxidized primarily at C2, C3 or C2/3 with the concomitant reduction of the flavin. A suitable electron acceptor then reoxidizes the reduced flavin. Whereas oxygen is a poor electron acceptor for PDH, several alternative acceptors, e.g., quinone compounds, naturally present during lignocellulose degradation, can be used. We have determined the 1.6-Å crystal structure of PDH from Agaricus meleagris. Interestingly, the flavin ring in PDH is modified by a covalent mono- or di-atomic species at the C(4a) position. Under normal conditions, PDH is not oxidized by oxygen; however, the related enzyme pyranose 2-oxidase (P2O) activates oxygen by a mechanism that proceeds via a covalent flavin C(4a)-hydroperoxide intermediate. Although the flavin C(4a) adduct is common in monooxygenases, it is unusual for flavoprotein oxidases, and it has been proposed that formation of the intermediate would be unfavorable in these oxidases. Thus, the flavin adduct in PDH not only shows that the adduct can be favorably accommodated in the active site, but also provides important details regarding the structural, spatial and physicochemical requirements for formation of this flavin intermediate in related oxidases. Extensive in silico modeling of carbohydrates in the PDH active site allowed us to rationalize the previously reported patterns of substrate specificity and regioselectivity. To evaluate the regioselectivity of D-glucose oxidation, reduction experiments were performed using fluorinated glucose. PDH was rapidly reduced by 3-fluorinated glucose, which has the C2 position accessible for oxidation, whereas 2-fluorinated glucose performed poorly (C3 accessible), indicating that the glucose C2 position is the primary site of attack.
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Affiliation(s)
- Tien Chye Tan
- School of Biotechnology, KTH Royal Institute of Technology, Stockholm, Sweden
- Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden
| | - Oliver Spadiut
- School of Biotechnology, KTH Royal Institute of Technology, Stockholm, Sweden
| | - Thanyaporn Wongnate
- Department of Biochemistry and Center of Excellence in Protein Structure and Function, Faculty of Science, Mahidol University, Bangkok, Thailand
| | - Jeerus Sucharitakul
- Department of Biochemistry, Faculty of Dentistry, Chulalongkorn University, Bangkok, Thailand
| | - Iris Krondorfer
- Food Biotechnology Laboratory, BOKU University of Natural Resources and Life Sciences, Vienna, Austria
| | - Christoph Sygmund
- Food Biotechnology Laboratory, BOKU University of Natural Resources and Life Sciences, Vienna, Austria
| | - Dietmar Haltrich
- Food Biotechnology Laboratory, BOKU University of Natural Resources and Life Sciences, Vienna, Austria
| | - Pimchai Chaiyen
- Department of Biochemistry and Center of Excellence in Protein Structure and Function, Faculty of Science, Mahidol University, Bangkok, Thailand
| | - Clemens K. Peterbauer
- Food Biotechnology Laboratory, BOKU University of Natural Resources and Life Sciences, Vienna, Austria
| | - Christina Divne
- School of Biotechnology, KTH Royal Institute of Technology, Stockholm, Sweden
- Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden
- * E-mail:
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