Characterisation of the First Archaeal Mannonate Dehydratase from Thermoplasma Acidophilum and Its Potential Role in the Catabolism of D-Mannose

catalysts

Article Characterisation of the First Archaeal Mannonate Dehydratase from Thermoplasma acidophilum and Its Potential Role in the Catabolism of D-Mannose

Dominik Kopp 1, Robert Willows 1,2 and Anwar Sunna 1,2,* 1 Department of Molecular Sciences, Macquarie University, Sydney, New South Wales 2109, Australia; [email protected] (D.K.); [email protected] (R.W.) 2 Biomolecular Discovery and Design Research Centre, Macquarie University, Sydney, New South Wales 2109, Australia * Correspondence: [email protected]; Tel.: +61-2-9850-4220

Received: 13 February 2019; Accepted: 24 February 2019; Published: 3 March 2019

Abstract: Mannonate dehydratases catalyse the dehydration reaction from mannonate to 2-keto-3-deoxygluconate as part of the hexuronic acid metabolism in bacteria. Bacterial mannonate dehydratases present in this gene cluster usually belong to the xylose isomerase-like superfamily, which have been the focus of structural, biochemical and physiological studies. Mannonate dehydratases from archaea have not been studied in detail. Here, we identified and characterised the first archaeal mannonate dehydratase (TaManD) from the thermoacidophilic archaeon Thermoplasma acidophilum. The recombinant TaManD enzyme was optimally active at 65 ◦C and showed high specificity towards D-mannonate and its lactone, D-mannono-1,4-lactone. The gene encoding for TaManD is located adjacent to a previously studied mannose-specific aldohexose dehydrogenase (AldT) in the genome of T. acidophilum. Using nuclear magnetic resonance (NMR) spectroscopy, we showed that the mannose-specific AldT produces the substrates for TaManD, demonstrating the possibility for an oxidative metabolism of mannose in T. acidophilum. Among previously studied mannonate dehydratases, TaManD showed closest homology to enzymes belonging to the xylose isomerase-like superfamily. Genetic analysis revealed that closely related mannonate dehydratases among archaea are not located in a hexuronate gene cluster like in bacteria, but next to putative aldohexose dehydrogenases, implying a different physiological role of mannonate dehydratases in those archaeal species.

Keywords: mannonate dehydratase; mannose metabolism; Thermoplasma acidophilum; mannono-1,4-lactone; 2-keto-3-deoxygluconate; aldohexose dehydrogenase

1. Introduction Mannonate dehydratases (EC 4.2.1.8) catalyse the conversion of mannonate to 2-keto-3-deoxygluconate (KDG) and have been studied as part of the hexuronic acid gene cluster in several organisms, such as Escherichia coli, Bacillus stearothermophilus, Bacillus subtilis and Erwinia chrysanthemi [1–4]. The hexuronate gene cluster encodes enzymes involved in the metabolism of glucuronate and galacturonate [5]. Glucuronate is a common sugar acid present in glucuronoxylan, a constituent of plant cell walls which can serve as the only carbon source for growth of some bacteria [3,6,7]. Glucuronate is also present in the mucus layer of mammals, providing a carbon source for anaerobic gut bacteria, such as E. coli [8,9]. As part of the hexuronate metabolism, mannonate dehydratase converts mannonate to KDG (Figure1A). In E. coli, and some species of Erwinia, KDG is phosphorylated and cleaved into pyruvate and 3-phosphoglycerate, which can be further metabolised in the tricarboxylic acid cycle, or the Entner-Doudoroff pathway [4,5].

Catalysts 2019, 9, 234; doi:10.3390/catal9030234 www.mdpi.com/journal/catalysts Catalysts 2019, 9, x FOR PEER REVIEW 2 of 14 Catalysts 2019, 9, 234 2 of 14

Mannonate dehydratases are represented in different enzyme families, such as the xylose isomeraseMannonate-like superfamily dehydratases or are the represented enolase superfamily. in different Mannonate enzyme dehydratasesfamilies, such encoded as the xylose in the isomerase-likebacterial hexuronate superfamily gene clusters or the usually enolase belong superfamily. to the xylose Mannonate isomerase dehydratases-like superfamily. encoded Crystal in thestructures bacterial have hexuronate been solved gene for clusters xylose isomerase usually belong-like mannonate to the xylose dehydratases isomerase-like from superfamily.Streptococcus Crystalsuis, E. coli structures K12 and haveEnterococcus been solved faecalis forin native xylose form isomerase-like and in complex mannonate with Mn2+ dehydratases ions [10,11]. His311 from Streptococcusand Tyr325 suisin the, E. binding coli K12 pock andetEnterococcus were identified faecalis asin crucial native for form the and activity in complex of the withmannonate Mn2+ ionsdehydratases [10,11]. His311 from Gram and Tyr325-positive in the bacteria binding, such pocket as S. were suis. identifiedHowever, asin crucialGram-negative for the activity bacteria of (e.g. the mannonateE. coli K12) dehydratases, an additional from inserted Gram-positive sequence in bacteria, the binding such aspocketS. suis rende. However,red the in dehydratase Gram-negative less bacteriaactive [11]. (e.g., MembersE. coli K12), of thean additional enolase superfamily inserted sequence show a in conserved the binding structure pocket rendered and reaction the dehydratasemechanism, less but active differ [ in11 ]. their Members physiological of the enolase functions. superfamily Within their show conserved a conserved barrel structure structure, and reactionmannonate mechanism, dehydratases but differ of this in their family physiological share conserved functions. ligand Within-bind theiring conserved sites for Mg barrel2+, which structure, are mannonateessential for dehydratases the stabilisation of thisof the family enediolate share intermediate conserved ligand-binding [12]. Among mannonate sites for Mg dehydratases2+, which are of essentialthe enolase for superfamily, the stabilisation some of therepresentatives enediolate intermediate with diverse [ 12functions]. Among have mannonate been found dehydratases, which are ofinvolved the enolase in the superfamily, catabolism some of sugar representatives acids other with than diverse glucuronate functions or have galacturonate been found, [13]. which Several are involvedcrystal structures in the catabolism have been of sugar solved acids for other mannonate than glucuronate dehydratases or galacturonate from the enolase [13]. Several superfamily crystal, structuresincluding have enzymes been from solved Chromohalobacter for mannonate salexigens dehydratases and fromNovosphingobium the enolase superfamily, aromaticivorans including [12,14]. enzymesHowever, from to theChromohalobacter best of our knowledge, salexigens and no archaealNovosphingobium mannonate aromaticivorans dehydratase[12 has,14 ].been However, investigated to the bestso far. of our knowledge, no archaeal mannonate dehydratase has been investigated so far. Here,Here, wewe usedused genomicgenomic analysisanalysis toto identifyidentify thethe firstfirst functionalfunctional mannonatemannonate dehydratasedehydratase fromfrom thethe thermoacidophilicthermoacidophilic archaeon archaeonThermoplasma Thermoplasma acidophilum acidophilum(TaManD). (TaMan The archaealD). The mannonate archaeal dehydratase mannonate wasdehydratase recombinantly-expressed was recombinantly in E.- coliexpressed, purified in and E. functionally coli, purified characterised. and functionally TaManD characterised. showed high aminoTaManD acid showed sequence high identity amino toacid bacterial sequence mannonate identity to dehydratases bacterial mannonate from the dehydratases xylose isomerase-like from the superfamily.xylose isomerase In the-like genome superfamily. of T. acidophilum In the genome, the geneof T. acidophilum encoding for, the TaManD gene encoding is located for adjacent TaManD to is anlocated aldohexose adjacent dehydrogenase to an aldohexose (AldT), dehydrogenase which has been (AldT), shown which previously has been to catalyse shown the previously oxidation to ofcatalyse mannose the to oxidation mannonate of with mannose high to specificity mannonate [15, 16 with]. However, high specificity the physiological [15,16]. However, significance the ofphysiological the oxidation sig andnificance its products of the were oxidation not investigated and its products further. were We identified not investigated the products further. of We an AldT-mediatedidentified the products oxidation of of an D-mannose AldT-mediated using NMRoxidation spectroscopy of D-mannose and confirmed using NMR that spectroscopy TaManD is able and toconfirmed convert the that products TaManD to is KDG. able Thisto convert demonstrates the products that in to principle, KDG. This a mannosedemonstrates metabolism that in principle, based on AldTa mannose and ManD metabolism is possible based in T.on acidophilum AldT and ManD(Figure is1 B).possible in T. acidophilum (Figure 1B).

UxaC UxuB UxuA KdgK glucuronate fructuronate mannonate KDG KDPG galucturonate tagaturonate altronate UxaC UxaB UxaA B CCM

AldT D-mannonate UxuA KDGA

D-mannose KDG CCM

D-mannono- 1,4-lactone

FigureFigure 1.1.Hexuronate Hexuronate metabolism metabolism in inE. E. coli coliand and possible possible mannose mannose catabolism catabolism in T. in acidophilum T. acidophilum.(A) Role. (A) ofRoleuxuA ofmannnonate uxuA mannnonate dehdyratase dehdyratase in dissimilation in dissimilation of hexuronates of hexuronates in E. coli adapted in E. from coli Peekhausadapted from and ConwayPeekhaus [8 ]. and (B) Role Conway of uxuA [8].mannonate (B) Role dehydratase of uxuA mannonate in a possible dehydratase mannose metabolism in a possible in T. acidophilum mannose. UxaC:metabolism hexuronate in T. acidophilum isomerase, UxuB:. UxaC: mannonate hexuronate oxidoreductase, isomerase, UxuB: UxaB: mannonate altronate oxidoreductase,oxidoreductase, UxuA: UxaB: mannonatealtronate oxidoreductase, dehydratase, UxaA: UxuA: altronate mannonate dehydratase, dehydratase, KdgK: 2-keto-3-deoxygluconateUxaA: altronate dehydratase, kinase, KdgK: KDG: 2-keto-3-deoxygluconate,2-keto-3-deoxygluconate KDPG: 2-keto-3-deoxy-6-phosphogluconate, kinase, KDG: 2-keto AldT:-3-deoxygluconate, aldose dehydrogenase, KDGA: KDPG: 2-keto-3-deoxygluconate2-keto-3-deoxy-6-phosphogluconate, aldolase, CCM: AldT: central aldose carbon dehydrogenase, metabolism. KDGA: 2-keto-3-deoxygluconate aldolase, CCM: central carbon metabolism.

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2. Results and Discussion

2.1. Screening for a Functional Mannonate Dehydratase in T. acidophilum Based on the previously identified mannose-specific AldT, we searched the genomic neighbourhood of its gene locus (Ta0754) for enzymes which could display activity towards mannonate. In the proximity of Ta0754, several genes are located that encode for hypothetical dehydratases, suggesting their potential to convert mannonate to KDG (Table S1). We identified the protein product of the Ta0753 gene as a functional mannonate dehydratase from T. acidophilum by heterologous expression in E. coli, and compared its properties and genomic context to other previously studied mannonate dehydratases. Among previously characterised mannonate dehydratases, TaManD shares highest protein sequence identity (31.5%) with enzymes in the xylose isomerase-like superfamily, such as the mannonate dehydratase from S. suis [10]. Key amino acid residues for substrate binding (His311 and Tyr325) and binding of the cofactor Mn2+ (Asp310, Cys237, His199 and His266) in S. suis are conserved in the amino acid sequence of TaManD. A much higher amino acid sequence identity (62.9%) is shared between TaManD and putative mannonate dehydratases from closely related archaeal species, Ferroplasma acidarmanus and Ferroplasma acidiphilum, suggesting an archaeal clade of mannonate dehydratases. Phylogenetic analysis showed that mannonate dehydratases from T. acidophilum, F. acidarmanus and F. acidiphilum are more closely related to bacterial mannonate dehydratases from the xylose isomerase-like superfamily than to bacterial or archaeal mannonate dehydratases of the enolase superfamily (Figure2). Mannonate dehydratases of the enolase superfamily have a substantially different structure from those in the xylose isomerase-like superfamily, and therefore, are only distantly related to TaManD. Despite fulfilling a similar function, the dihydroxy-acid dehydratase from S. solfataricus, which belongs to the IlvD/EDD superfamily, is rather unrelated to the xylose isomerase-like and enolase superfamilies [17]. For bacterial xylose isomerase-like mannonate dehydratases, a physiological function of the enzyme in the catabolism of hexuronates has been demonstrated [18,19]. Although TaManD is annotated as uxuA mannonate dehydratase, which implies a role in the metabolism of hexuronates, it is not present in a classical hexuronate gene cluster known from bacteria such as E. coli, B. subtilis or B. stearothermophilus (Figure3)[ 2,3]. In contrast, the gene encoding for TaManD is located adjacent to AldT in the genome of T. acidophilum. Similarly, the two closely related putative mannonate dehydratases in F. acidarmanus and F. acidiphilum are also located adjacent to putative aldohexose dehydrogenases. Therefore, a different physiological role can be proposed for these mannonate dehydratases. In the following, we functionally characterise TaManD and show that AldT is able to produce the substrate needed for a subsequent conversion to KDG mediated by TaManD. Catalysts 2019, 9, x FOR PEER REVIEW 3 of 14

2. Results and Discussion

2.1. Screening for a functional mannonate dehydratase in T. acidophilum Based on the previously identified mannose-specific AldT, we searched the genomic neighbourhood of its gene locus (Ta0754) for enzymes which could display activity towards mannonate. In the proximity of Ta0754, several genes are located that encode for hypothetical dehydratases, suggesting their potential to convert mannonate to KDG (Table S1). We identified the protein product of the Ta0753 gene as a functional mannonate dehydratase from T. acidophilum by heterologous expression in E. coli, and compared its properties and genomic context to other previously studied mannonate dehydratases. Among previously characterised mannonate dehydratases, TaManD shares highest protein sequence identity (31.5%) with enzymes in the xylose isomerase-like superfamily, such as the mannonate dehydratase from S. suis [10]. Key amino acid residues for substrate binding (His311 and Tyr325) and binding of the cofactor Mn2+ (Asp310, Cys237, His199 and His266) in S. suis are conserved in the amino acid sequence of TaManD. A much higher amino acid sequence identity (62.9%) is shared between TaManD and putative mannonate dehydratases from closely related Catalysts 2019archaeal, 9, 234 species, Ferroplasma acidarmanus and Ferroplasma acidiphilum, suggesting an archaeal clade of 4 of 14 mannonate dehydratases.

Catalysts 2019, 9, x FOR PEER REVIEW 4 of 14

crescentus/Caulobacter vibroides: Q9AAR4, Novosphingobium aromaticivorans: A4XF23, Chromohalobacter salexigens: Q1QT89.

Phylogenetic analysis showed that mannonate dehydratases from T. acidophilum, F. acidarmanus and F. acidiphilum are more closely related to bacterial mannonate dehydratases from the xylose isomerase-like superfamily than to bacterial or archaeal mannonate dehydratases of the enolase superfamily (Figure 2). Mannonate dehydratases of the enolase superfamily have a substantially Figure 2.FigurePhylogenetic 2. Phylogenetic relationship relationship of different of different putative putative and and confirmed confirmed dehydratases dehydratases belonging belonging to to different different structure from those in the xylose isomerase-like superfamily, and therefore, are only enzymedifferent families. enzyme Characterised families. Characterised enzymes are enzymes underlined. are underlined. Evolutionary Evolutionary analyses analyses were conducted were in distantlyconducte relatedd into MEGA TaManD. 7. 20 protein Despite sequences fulfilling were a alignedsimilar using function, MUSCLE. the Thedihydroxy phylogenetic-acid tree dehydratase was MEGA 7. 20 protein sequences were aligned using MUSCLE. The phylogenetic tree was inferred using from S.inferred solfataricus using, thewhich neighbour belongs-joining to tmethod.he IlvD/EDD The scale superfamily, bar indicates an is evolutionary rather unrelated distance toof 0.20the xylose theisomerase neighbour-joiningnucleotide-like and per enolase position method. superfamilies in The the scale sequence. bar [17]. indicates The number an evolutionary next to the nod distancees represent of 0.20 bootstrap nucleotide per positionForconfidence in bacterial the sequence. values xylose estimated The isomerase number from- 500 nextlike replicates. mannonateto the nodes Protein dehydratases, represent sequences bootstrap were a retrieved physiological confidence from the values function Uniprot estimated of the fromenzyme 500database in replicates. the andcatabolism Protein their entry sequences of hexuronates codes are were as retrieved hasfollows. been Ferroplasma from demonstra the Uniprot acidarmanusted [18,19]. database: S0AL33, Although and Ferroplasma their TaManD entry codes is areannotated as follows.acidiphilum as uxuAFerroplasma: A0A1V0N416,mannonate acidarmanus dehydratase,Thermoplasma: S0AL33, which acidFerroplasmaophilum implies: Q9HK52, a acidiphilum role inPicrophilus the: metabolism A0A1V0N416, torridus: of Q6L2R9, Thermoplasmahexuronates, acidophilumit is notVulcanisaeta present: Q9HK52, in moutnovskiaa classicalPicrophilus: hexuronate F0QYL3, torridus: Q6L2R9,Caldivirga gene cluster Vulcanisaetasp. JCHS known 4: moutnovskia from A0A101XEY7, bacteria: F0QYL3, SulfolobussuchCaldivirga as E.sp. coli A20:,sp. B. JCHSsubtilis 4: A0A1C8ZTN0, Escherichia coli K12: P24215, Erwinia chrysanthemi/Dickeya dadantii: E0SEP1, A0A101XEY7,or B. stearothermophilSulfolobusus (Figsp. A20:ure 3) A0A1C8ZTN0, [2,3]. In contrast,Escherichia the gene coli encodingK12: P24215, for TaManDErwinia chrysanthemi is located adjacent/Dickeya Streptococcus suis: A0A142UME2, Enterococcus faecalis: Q82ZC9, Thermotoga maritima: Q9WXS4, dadantiito AldT: E0SEP1, in the genomeStreptococcus of T. suis acidophilum: A0A142UME2,. Similarly,Enterococcus the two faecalis closely: Q82ZC9, related putativeThermotoga mannonate maritima: Bacillus subtilis: O34346, Bacillus stearothermophilus: A0A087LHB3, Sulfolobus solfataricus DHAD: Q9WXS4,dehydratasesQ97UB2,Bacillus in Sulfol F. subtilis acidarmanusobus: acidocaldarius O34346, andBacillus DHAD:F. acidiphilum stearothermophilus Q4J860, Sulfolobusare also: A0A087LHB3, solfataricuslocated adjacent GNAD:Sulfolobus Q97U27, to putative solfataricus Caulobacter aldohexose DHAD: Q97UB2,dehydrogenases.Sulfolobus Therefore, acidocaldarius a differentDHAD: physiological Q4J860, Sulfolobus role cansolfataricus be proposedGNAD: for Q97U27, these mannonateCaulobacter crescentusdehydratases./Caulobacter In the following, vibroides: Q9AAR4,we functionallyNovosphingobium characterise aromaticivorans TaManD and: show A4XF23, thatChromohalobacter AldT is able to

salexigensproduce :the Q1QT89. substrate needed for a subsequent conversion to KDG mediated by TaManD.

FigureFigure 3. Schematic 3. Schematic maps maps of of uxuA uxuA mannonatemannonate dehydratases dehydratases and and their their genomic genomic neighbourhoods neighbourhoods in T. in T. acidophilumacidophilum, ,F. F. acidarmanusacidarmanus, ,F.F. acidiphilum acidiphilum andand B. stearothermophilusB. stearothermophilus. Genes. are Genes annotated are annotated in the NCBI in the NCBIdatabase database as as follows: follows: In In blue blue,, TaManDTaManD/uxuA/uxuA: T.: T. acidophilum acidophilum mannonatemannonate dehydratase/uxuA dehydratase/uxuA D-mannonateD-mannonate dehydratases; dehydratases in; in orange, orange,AldT AldT/AlDH/AlDH/GDHGDH: : aldohexose aldohexose dehydrogenase/aldose dehydrogenase/aldose dehydrogenase/glucose-1-dehydrogenase;dehydrogenase/glucose-1-dehydrogenase; in in green green,, TR: TR: transcriptional transcriptional regulator regulator;; in light in light grey, grey, annotatedannotated as hypothetical as hypothetical protein. protein. Note: Note: 750:750: FAA hydrolase hydrolase family family protein, protein, 752: 752: L-rhamnonate L-rhamnonate dehydratase, 756: mandelate racemase/muconate lactonizing enzyme family protein, 460: dehydratase, 756: mandelate racemase/muconate lactonizing enzyme family protein, 460: transposase, 485: acetyl-coenzyme A synthetase, 1022: acetate-CoA ligase, xynB: β-xylosidase, kdgK: transposase, 485: acetyl-coenzyme A synthetase, 1022: acetate-CoA ligase, xynB: β-xylosidase, kdgK: 2-keto-3-deoxygluconate kinase, kdgA: 2-keto-3-deoxy-6-phosphogluconate aldolase, uxaR: 2-keto-3-deoxygluconate kinase, kdgA: 2-keto-3-deoxy-6-phosphogluconate aldolase, uxaR: regulatory regulatory protein, uxaC: hexuronate isomerase, uxuB: mannonate oxidoreductase. protein, uxaC: hexuronate isomerase, uxuB: mannonate oxidoreductase. 2.2. Substrate conformation for TaManD activity Before acquiring enzyme kinetics and studying the characteristics of TaManD, we investigated which substrate conformation is encountered by TaManD under physiological conditions. In aqueous solutions, free sugar acids coexist with their lactone in an equilibrium, which is defined by the stability of the lactone, the temperature and the pH of the solution. Many lactones hydrolyse spontaneously in water, although several lactones, including D-arabinonolactone [20], L-rhamnonolactone [21] and D-xylonolactone [22], have been reported to be hydrolysed by

Catalysts 2019, 9, 234 5 of 14

2.2. Substrate Conformation for TaManD Activity Before acquiring enzyme kinetics and studying the characteristics of TaManD, we investigated which substrate conformation is encountered by TaManD under physiological conditions. In aqueous solutions, free sugar acids coexist with their lactone in an equilibrium, which is deﬁned by the stability of the lactone, the temperature and the pH of the solution. Many lactones hydrolyse spontaneously in water, although several lactones, including D-arabinonolactone [20], L-rhamnonolactone [21] and D-xylonolactone [22], have been reported to be hydrolysed by lactonases. Mannonate is able to form a lactone by covalent bond formation between carbon 1 and carbon 5 (δ-lactone) or carbon 1 and carbon 4 (γ-lactone) (Figure4A). The equilibrium of the two different envelope forms has been studied by NMR spectroscopy and it was found that the γ-lactone is strongly favoured over the δ-lactone [23]. However, these studies do not describe equilibria between lactone and free acid form in a physiological buffer. In a cellular environment mannonate can either be produced by a mannonate oxidoreductase (UxuB) as part of the catabolism of hexuronic acids or by an aldohexose dehydrogenase, such as AldT, in a hypothetical oxidative mannose catabolism (Figure1). In order to study the conformation of the substrate for TaManD in a physiological environment, we ﬁrst acquired decoupled 13C NMR spectra for D-mannono-1,4-lactone in physiological buffer at pH 7, in its lactone form (incubated with HCl) and after hydrolysis to the free sugar acid (incubated with NaOH) (Figure4B). The spectra obtained indicated that the lactone and the free sugar acid can be distinguished by their chemical shifts. In the free sugar acid form, carbon 6 yields a chemical shift at 63.51 ppm, whereas in D-mannono-1,4-lactone, carbon 6 displays a chemical shift at 63.02 ppm. D-mannono-1,4-lactone in buffer shows both chemical shifts, indicating that D-mannono-1,4-lactone and D-mannonate are present in an equilibrium at pH 7.

63.51 A B C6 OH OH O HO 63.02 OH OH O NaOH D-mannonate

HCl HO O O HO buffer

HO OH D-mannono-1,4-lactone ppm 180 80 75 70 65 60 C

D-[1,6-13C ]mannose 2 63.51 63.02

175 95 90 65 60

13 D-[1,6- C2]mannose oxidation

175 95 90ppm 65 60

Figure 4. Analysis of D-mannono-1,4-lactone under different conditions and products of AldT-mediated oxidation of isotope labelled D-mannose. (A) Chemical structures of the free sugar acid, D-mannonate, and its lactone, D-mannono-1,4-lactone. Carbon 6, which was used to differentiate the two different substrate forms in NMR spectra, is circled for both structures. (B) Decoupled 1D 13C NMR spectra of D-mannono-1,4-lactone after incubation in 1 M NaOH, 1 M HCl and in 0.1 M sodium phosphate (NaP) buffer pH 7. Chemical shift for carbon 6 is shown by a dashed rectangle. Characteristic chemical shifts for the lactone (63.02 ppm) and for the free sugar acid (63.51 ppm) are indicated. (C) Decoupled 1D 13C NMR spectra before (top) and after (bottom) oxidation of labelled D-mannose with AldT. Chemical shifts of carbon 6 for lactone and free sugar acid are indicated. Catalysts 2019, 9, 234 6 of 14

13 Next, we oxidised isotope-labelled D-[1,6- C2]mannose using recombinantly-expressed AldT and acquired 1D 13C NMR spectra of the reaction product. The chemical shifts for carbon 6 of mannono-1,4-lactoneCatalysts 2019, 9, x FOR PEER and REVIEW D-mannonate were used to analyse the reaction products of the mannose6 of 14 oxidation with AldT, as these were close together but well resolved and will have similar relaxation oxidation with AldT, as these were close together but well resolved and will13 have similar relaxation rates in both forms. The spectrum after partial AldT oxidation of D-[1,6- C2]mannose (Figure4C) 13 showsrates thatin both equal forms. quantities The spectrum of D-mannono-1,4-lactone after partial AldT oxidation and D-mannonate of D-[1,6- C2]mannose are produced (Figure for 4C) the subsequentshows that dehydration equal quantities by TaManD. of D-mannono-1,4-lactone and D-mannonate are produced for the subsequent dehydration by TaManD. 2.3. Expression and Purification of TaManD 2.3. Expression and purification of TaManD For the further biochemical characterisation, TaManD was expressed together with a tobacco etch For the further biochemical characterisation, TaManD was expressed together with a tobacco virus (TEV) sequence, which allowed the proteolytic cleavage of the His-tag after the first metal affinity etch virus (TEV) sequence, which allowed the proteolytic cleavage of the His-tag after the first metal purification. In order to study the effect of bivalent metal cofactors on enzyme activity, cleavage of affinity purification. In order to study the effect of bivalent metal cofactors2+ on2+ enzyme activity, the His-tag from the recombinant enzyme was necessary, since cations (Ni , Mg ) might influence cleavage of the His-tag from the recombinant enzyme was necessary, since cations (Ni2+, Mg2+) might enzyme activity assays of a His-tagged enzyme. After His-tag cleavage, the recombinant enzyme influence enzyme activity assays of a His-tagged enzyme. After His-tag cleavage, the recombinant showedenzyme a single showed band a single on sodium band dodecyl on sodium sulfate dodecyl polyacrylamide sulfate polyacrylamide gel electrophoresis gel electrophoresis (SDS-PAGE) with(SDS an- apparentPAGE) with mass an of apparent 38 kDa (Figure mass S1).of 38 Peptide kDa (Fig massure fingerprinting S1). Peptide was mass performed fingerprinting with liquidwas chromatography-electrosprayperformed with liquid chromatography ionisation-tandem-electrospray mass spectrometryionisation-tandem (LC ESI mass MS/MS) spectrometry and the (LC National ESI CenterMS/MS for)Biotechnology and the National Information Center for (NCBI)Biotechnology database Information searches using (NCBI the) database mascot searches software using confirmed the themascot identity software of the purified confirmed protein the identity as uxuA of mannonate the purified dehydratase protein as uxuAfrom T. mannonate acidophilum dehydratase. In addition, thefrom apparent T. acidophilum native molecular. In addition, mass the of theapparent protein native was estimatedmolecular frommass sizeof the exclusion protein chromatographywas estimated to befrom 225 size kDa, exclusion indicating chromatography a hexameric structureto be 225 kDa, of the indicating protein. a hexameric structure of the protein.

2.4.2.4. Enzyme Enzyme Characterisation characterisation InitialInitial tests tests showed showed that that TaManD TaManD lost lost almost almost its its complete complete activity activity after after purification, purification suggesting, suggesting that essentialthat essential cofactors cofactors for activity for activity were removed were removed during during the purification the purification process. process. Accordingly, Accordingly, the effect the of severaleffect additives of several was additives tested for wastheir influencetested for on their the activity influence of TaManD on the withactivity D-mannono-1,4-lactone. of TaManD with TaManDD-mannono showed-1,4- stronglylactone. enhanced TaManD activityshowed in strongly the presence enhanced of β -mercaptoethanol activity in the and presence the metal of ionsβ- Mnmercaptoethanol2+, Ni2+, Mg2+ and, Ca 2+theand metal Co2+ ions(Figure Mn2+5,). Ni The2+, highestMg2+, Ca activation2+ and Co (100%)2+ (Figure was 5). observed The highest in the presenceactivation of β -mercaptoethanol, (100%) was observed which in was the significantlypresence of β higher-mercaptoethanol, than the activating which bivalent was significantly cation Ni 2+ 2+ (71.5%higher of maximumthan the activating enzyme bivalent activity, cation p-value: Ni 0.0022). (71.5% of Mn maximum2+, Mg2+ enzymeand Co activity,2+ also activated p-value: 0.0022). TaManD 2+ 2+ 2+ andMn resulted, Mg in and 50–60% Co also increase activated in maximum TaManD activity and resulted when comparedin 50–60% toincrease the TaManD in maximum control activity (without when compared to the TaManD control (without any additive). No significant difference in activity any additive). No significant difference in activity was observed in the presence of Fe2+, Zn2+, Cu2+, was observed in the presence of Fe2+, Zn2+, Cu2+, EDTA, DTT and glutathione. EDTA, DTT and glutathione.

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l e r 20

0 l + + + + + + + + o 2 s 2 2 2 2 2 2 2 A T E e e tr y i T T M in n n e c n n N g a u o D - e io F M Z M C C C D t h o + + E s t c 2 y ta e c lu F g

Figure 5. Effect of metal ions, chelating and reducing agents on the activity of TaManD. Puriﬁed enzyme Figure 5. Effect of metal ions, chelating and reducing agents on the activity of TaManD. Purified (0.6 µg) was pre-incubated for 1 h with 1 mM of each additive in 50 mM HEPES pH 7. Activity was then enzyme (0.6 µg) was pre-incubated for 1 h with 1 mM of each additive in 50 mM HEPES pH 7. determined in reactions containing 10 mM D-mannono-1,4-lactone in 50 mM HEPES pH 7 incubated Activity was then determined in reactions containing 10 mM D-mannono-1,4-lactone in 50 mM for 1 h at 55 ◦C before analysis with the semicarbazide assay. Activity is expressed in relation to the HEPES pH 7 incubated for 1 h at 55 ˚C before analysis with the semicarbazide assay. Activity is maximum enzyme activity. Fe2+ + cys: Fe2+ was prepared with 1 mM cysteine. DTT: dithioerythritol, expressed in relation to the maximum enzyme activity. Fe2+ + cys: Fe2+ was prepared with 1 mM EDTA: ethylendiaminetetraacetate, β-ME: β-mercaptoethanol. cysteine. DTT: dithioerythritol, EDTA: ethylendiaminetetraacetate, β-ME: β-mercaptoethanol.

TaManD was activated by similar bivalent ions and reducing factor, as previously studied bacterial mannonate dehydratases from the xylose isomerase-like superfamily. However, none of

Catalysts 2019, 9, 234 7 of 14

Catalysts 2019, 9, x FOR PEER REVIEW 7 of 14 TaManD was activated by similar bivalent ions and reducing factor, as previously studied bacterial mannonatethe bacterial dehydratases mannonate dehydratases from the xylose described isomerase-like so far have superfamily. been shown However, to have none an ofequally the bacterial strong mannonateactivation with dehydratases both metal described ions, Co2+ so and far Ni have2+, as been observed shown with to have TaManD. an equally Early stronginvestigations activation of withmannonate both metal dehydratases ions, Co2+ focusedand Ni2+ on, as E. observed coli and with showed TaManD. differ Earlyent strengths investigations in activation of mannonate after dehydratasesincubation with focused bivalent on E. coli cationsand showed [1,24– different26]. In strengths summary, in activation most studies after incubation showed with that bivalentβ-mercaptoethanol, cations [1,24 – Fe262+]. and In summary, Mn2+ resulted most studies in highest showed enzyme that β -mercaptoethanol, activity (90–100% Fe of2+ maximumand Mn2+ resultedenzyme inactivity). highest Incubation enzyme activity with other (90–100% bivalent of maximum cations resulted enzyme in activity). less activity Incubation compared with to other the bivalentenzyme's cations maximum resulted activity in less (Co activity2+: 27–80%, compared Ni2+: 5 to–40% the enzyme’sand Zn2+: maximum5–25%, depending activity (Coon 2+the: 27–80%,study). NiMore2+: 5–40%recently, and crystal Zn2+ :structures 5–25%, depending of xylose isomerase on the study).-like Moremannonate recently, dehydratases crystal structures from S. of suis xylose and isomerase-likeE. coli have been mannonate solved and dehydratases revealed the presence from S. suisof primarilyand E. coli Mnhave2+ and been lower solved amounts and revealedof Mg2+, Ni the2+ presenceand Zn2+ ofin primarilytheir binding Mn 2+sitesand [10,11]. lower amounts of Mg2+, Ni2+ and Zn2+ in their binding sites [10,11]. T. acidophilumacidophilum was originally isolated from a hot and acidic environmentenvironment and accordingly this archaeon displays optimal temperatu temperaturere and pH for growth at 59 °◦C and pH 1 1–2,–2, respectively [27]. [27]. The puriﬁedpurified TaManD waswas activeactive betweenbetween 3535 ◦°C andand 70 ◦°C, with an optimal temperaturetemperature for activity at 65 ◦°C (Figure(Figure6 6A).A). Thermostability Thermostability of of the the enzyme enzyme in in the the absence absence of of substrate substrate was was studied studied at at temperaturestemperatures between 5555 ◦°C and 95 ◦°C (Fig (Figureure6 6B).B). TheThe enzymeenzyme retainsretains itsits fullfull activityactivity atat 5555 ◦°C for at least an hour, whereaswhereas completecomplete inactivationinactivation was observed at 75 ◦°C, 85 ◦°C and 95 °◦C within 90 min, 60 min and 1515 min,min, respectively.respectively. The difference in the relatively high optimal temperature of the enzyme to to the the comparably comparably low low thermostability thermostability indicated indicated that the that enzyme the is enzyme more prone is more to inactivation prone to ininactivation the absence in of the substrate. absence Other of substrate. enzymes Other obtained enzymes from this obtained organism from have this been organism shown have to display been optimalshown to temperatures display optimal in the tempe rangeratures of 55 ◦inC the to 70range◦C[ 15of ,5528 –°30C ].to 70 °C [15,28–30].

A B

150 )

) 100

(

y t

t 100 i

i 75

i t

t 55˚C

65˚C e

e 50

i t

t 50 75˚C

l e

e 25 r r 85˚C 95˚C 0 0 30 40 50 60 70 0 20 40 60 80 100 temperature (oC) time (min)

Figure 6. Effect of temperature on TaManD activity. (A) Optimum temperature of TaManD. (FigureB) Thermostability 6. Effect of oftemperature purified TaManD on TaManD enzyme activity. (1 µg) incubated (A) Optimum over 90 temperature min at various of temperatures.TaManD. (B) Thermostability of purified TaManD enzyme (1 µg) incubated over 90 min at various temperatures. Following the identification of the crucial metal cofactors required for TaManD activity and determinationFollowing of the its id optimalentification reaction of the temperature, crucial metal we acquired cofactors enzyme required kinetic for TaManD data for activitythe purified and enzyme.determination Considering of its optimal the fact reaction that under temperature, physiological we acquired conditions enzyme an equilibriumkinetic data offor boththe purified lactone andenzyme. free Considering sugar acid exists, the fact the that kinetic under data physiological was acquired conditions in reactions an equilibrium with substrate of both provided lactone and in thesefree sugar two differentacid exists, forms. the kinetic The substrate data was was acquired either hydrolysedin reactions withwith NaOHsubstrate to obtainprovided free in sugar these acid,two D-mannonate,different forms. or the The lactone substrate was prepared was either in buffer, hydrolysed equaling with an equilibrium NaOH to obtain of lactone free and sugar free sugar acid, acid.D-mannonate, TaManD displayedor the lactone a higher was activity prepared in reactionsin buffer, with equaling D-mannonate an equilibrium compared of tolactone reactions and withfree D-mannono-1,4-lactonesugar acid. TaManD displayed prepared in a buffer higher at pHactivity 7 (Table in 1 reactions). The maximal with velocity D-mannonate of the reaction, compared V max to, withreactions D-mannonate with D-mannono as substrate-1,4-lactone was slightly prepar highered in buffer (24–27%) at pH compared 7 (Table to 1). reactions The maximal with thevelocity lactone of preparedthe reaction, in buffer Vmax (p-value:, with D 0.06),-mannonate whereas as no substrate difference was in the slightly affinity betweenhigher (24 the–27% two)different compared forms to ofreactions the substrate with the could lactone be observed prepared (Table in buffer1, Figure (p-value: S2). 0.06), whereas no difference in the affinity betweenThe the pH two optimum different of TaManDforms of wasthe substrate dependent cou onld thebe observed form of the (Table substrate. 1, Figure TaManD S2). displayed highest activity between pH 5 and 7 with mannonate, while with D-mannono-1,4-lactone in buffer, the enzymeTable 1. TaManD displayed kinetic overall data lowerwith D activities-mannonate with (prepared an apparent by hydrolysis optimum with NaOH) at pH and 7 (Figure with 7). UnlikeD- previouslymannono-1,4 studied-lactone enzymesprepared in from bufferT. acidophilum(pH 7). Non-(e.g.,linear AldT), fitting was TaManD performed did not (Fig retainure S2 maximum). activity above pH 7 with both substrates [15]. substrate Km (mM) Vmax (U/mg) kcat (s−1) kcat/Km (mM−1 s−1) D-mannonate 5.37 ± 0.90 2.39 ± 0.11 1.64 0.30 D-mannono-1,4-lactone in buffer 4.90 ± 0.53 1.90 ± 0.06 1.33 0.26

Catalysts 2019, 9, x FOR PEER REVIEW 8 of 14

The pH optimum of TaManD was dependent on the form of the substrate. TaManD displayed highest activity between pH 5 and 7 with mannonate, while with D-mannono-1,4-lactone in buffer, the enzyme displayed overall lower activities with an apparent optimum at pH 7 (Figure 7). Unlike previously studied enzymes from T. acidophilum (e.g. AldT), TaManD did not retain maximum activity above pH 7 with both substrates [15].

CatalystsBased2019, 9 on, 234 the observations from experiments with different forms of substrate, we assume8 ofthat 14 D-mannonate is either the only form, or at least more accessible to TaManD than its lactone. At low pH, TaManD shows low activity with D-mannono-1,4-lactone, supposedly because the lactone does not Table hydrolyse 1. TaManD to the kinetic free data acid. with InD-mannonate contrast, (prepared if D-mannono by hydrolysis-1,4-lactone with NaOH) was hyd androlysed with to D-mannonateD-mannono-1,4-lactone and used as prepared substrate in bufferat low (pH pH, 7). high Non-linear enzyme ﬁtting activity was should performed be (Figureobserved. S2). Similarly, the difference in Vmax for D-mannonate and D-mannono-1,4-lactone in− 1buffer can be explained−1 −1 by Substrate Km (mM) Vmax (U/mg) kcat (s ) kcat/Km (mM s ) slow hydrolysis of the lactone to the free acid. Reactions with hydrolysed D-mannonate occur faster D-mannonate 5.37 ± 0.90 2.39 ± 0.11 1.64 0.30 comparedD-mannono-1,4-lactone to the lactone inin buffer buffer, since 4.90 more± 0.53 substrate 1.90 is± readily0.06 available. 1.33 0.26

125 mannonate lactone in buffer

) 100

(

y t

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l e r 25

0 4 6 8 10 12 pH FigureFigure 7. 7.The The pH pH optima optima of of reactions reactions with with D-mannono-1,4-lactone D-mannono-1,4-lactone prepared prepared in bufferin buffer pH pH 7 (lactone 7 (lactone in buffer)in buffer) and with and D-mannonate with D-mannonate (prepared (prepared by hydrolysis by hydrolysis with NaOH). with Activity NaOH). values Activity are expressed values asare relativeexpressed activity as relative (%) normalised activity (%) to thenormalised highest overallto the highest activity overall observed activity in the observed assay. in the assay.

2.5. BasedProduct on identification the observations and substrate from experiments specificity with different forms of substrate, we assume that D-mannonate is either the only form, or at least more accessible to TaManD than its lactone. At low pH, TaManDIn order shows to identify low activity the product with D-mannono-1,4-lactone,of the TaManD reaction with supposedly mannonate, because reactions the lactone containing does 2+ notthe hydrolyse substrate to andthe free Co acid. were In contrast, analysed if D-mannono-1,4-lactone at several time points was by hydrolysed high-performance to D-mannonate liquid andchromatography used as substrate (HPLC at low) (Fig pH,ure high 8A). enzyme Although activity a complete should be separation observed. of Similarly, the product the difference KDG and mannonate could not be achieved on different organic acid columns, an increase of KDG was in Vmax for D-mannonate and D-mannono-1,4-lactone in buffer can be explained by slow hydrolysis ofobserved. the lactone In to order the free to acid.test for Reactions substrate with promiscuity, hydrolysed D-mannonate11 different sugar occur acids faster (Fig comparedure 8B) to were the lactoneincubated in buffer, with purified since more TaManD substrate and is tested readily for available. the formation of 2-keto-3-deoxy analogues using the semicarbazide assay. D-mannonate and D-mannono-1,4-lactone were the only substrates that 2.5.showed Product a Identification positive reaction and Substrate after incubation Specificity for 16 h. This is in contrast to other dehydratases, including members of the enolase superfamily and the IlvD/EDD superfamily, such as the In order to identify the product of the TaManD reaction with mannonate, reactions containing the L-fuconate dehydratase from Xanthomonas campestris [31] or the dihydroxy-acid dehydratase from substrate and Co2+ were analysed at several time points by high-performance liquid chromatography Sulfolobus solfataricus [17,32], which have been shown to be active with a multitude of different sugar (HPLC) (Figure8A). Although a complete separation of the product KDG and mannonate could not acids. be achieved on different organic acid columns, an increase of KDG was observed. In order to test for substrate promiscuity, 11 different sugar acids (Figure8B) were incubated with purified TaManD and tested for the formation of 2-keto-3-deoxy analogues using the semicarbazide assay. D-mannonate and D-mannono-1,4-lactone were the only substrates that showed a positive reaction after incubation for 16 h. This is in contrast to other dehydratases, including members of the enolase superfamily and the IlvD/EDD superfamily, such as the L-fuconate dehydratase from Xanthomonas campestris [31] or the dihydroxy-acid dehydratase from Sulfolobus solfataricus [17,32], which have been shown to be active with a multitude of different sugar acids.

Catalysts 2019, 9, 234 9 of 14 Catalysts 2019, 9, x FOR PEER REVIEW 9 of 14

A B

tested sugar acids relative activity 6000 D-mannonate 100% mannonate 5000 D-mannono-1,4-lactone 79% KDG 30min D-gluconic acid sodium salt n.d. 4000 60min

U D-galactono-1,4-lactone n.d.

I 120 min R 3000 D-glucuronic acid n.d. D-galacturonic acid sodium salt n.d. 2000 L-gulonic acid- -lactone n.d.

1000 D-arabinoic acid sodium salt n.d.

D-fuconic acid lithium salt n.d. 0 11 12 13 14 15 L-fuconic acid sodium salt n.d. retention time (min) L-rhamnonic acid lithium salt n.d.

Figure 8. Substrate specificity and product analysis of reactions with TaManD. (A) Reactions with FigureTaManD 8. Substrate and 10 mM spe mannonatecificity and product analysed analysis by HPLC of reactions equipped with with TaManD a refractive. (A) Reactions index detector. with TaManDStandards: and 10 mM 10 mM mannonate mannonate (dashed analysed black line), by HPLC 2.5 mM equipped KDG (dashed with grey a refractive line). Reactions: index detector. stopped Standards:after 30 min 10 (red mM line), mannonate stopped after (dashed 60 min black (blue line), line), stopped 2.5 mM after KDG 120 (dashed min (green grey line). line). RIU: Reactions: refractive stopindexped units. after (B30) Listmin of(red sugar line), acids stopped that wereafter tested60 min as (blue substrates line), stopped for TaManD. after 120 Activity min (green is expressed line). RIU:relative refractive (%) to the index maximal units. activity(B) List obtainedof sugar inacids the assay;that were n.d.: tested no signal as substrates detected afterfor TaManD. 16 h of incubation. Activity isLactones expressed except relative for D-mannono-1,4-lactone(%) to the maximal activity were ob hydrolysedtained in the by assay incubation; n.d.: no in 1signal M NaOH detected for 1after h at 16room h of temperatureincubation. Lactones and then except diluted for in D 50-mannono mM HEPES-1,4- pHlactone 7 to obtainwere hydrolysed free sugar acids.by incubation All sugar in acids 1 M NaOHwere tested for 1 ath aat concentration room temperature of 10 mM and in then duplicate diluted experiments. in 50 mM HEPES pH 7 to obtain free sugar acids. All sugar acids were tested at a concentration of 10 mM in duplicate experiments. 3. Materials and Methods 3. Materials and Methods 3.1. Cloning, Expression and Purification of Enzymes 3.1. Cloning,All plasmids expression were and constructed purification by of amplificationenzymes of genes from T. acidophilum DSM 1728 genomic DNAAll (DSMZ, plasmids Braunschweig, were constructed Germany) by amplification using PCR, of restriction genes from and T. acidophilum ligation into DSM vectors 1728 according genomic DNAto standard (DSMZ, protocols Braunschweig, [33]. Primer Germany) pairs using and restrictionPCR, restriction enzymes and usedligation for into the constructionvectors according of each to α standardexpression protocols plasmid [3 are3]. summarised Primer pairs in and Table restriction S2. E. coli enzymestrains used-select for (Bioline, the construction Sydney, Australia) of each expressionwas used inplasmid all initial are cloning summarised procedures. in Table The S2. expression E. coli strain plasmid α-select pProEX (Bioline, HTa-Ta0753 Sydney, Australia) was then wasused used to transform in all initial BL21-CodonPlus cloning procedures. (DE3)-RIL The competent expression cells plasmid (Agilent pProEX Technologies, HTa-Ta0753 Santa Clara,was then CA, usedUSA). to Plasmid transform pETDuet-1-AldT BL21-CodonPlus was (DE3) used- toRIL transform competent BL21 cells (DE3) (Agilent cells (NEB,Technologies, Ipswitch, Santa Burlington, Clara, CAMA,, USA). USA). PlasmidBL21-CodonPlus pETDuet (DE3)-RIL-1-AldT was strains used containing to transform the respective BL21 (DE3) expression cells (NEB, plasmid Ipswitch, were µ Burlington,grown in 500 MA, mL USA). Luria–Bertani BL21-CodonPlus medium (DE3) (LB) supplemented-RIL strains contai withning carbenicillin the respective (100 expressiong/ml) and µ ◦ plasmidchloramphenicol were grown (35 ing/ml) 500 at ml 37 LuriaC to–Bertani an OD600 mediumof 0.6 before (LB) supplemented induction was performedwith carbenicillin with 0.4 (100 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) at 20 ◦C for 16–18 h. The BL21 (DE3) cells carrying µg/ml) and chloramphenicol (35 µg/ml) at 37 ˚C to an OD600 of 0.6 before induction was performed withthe expression 0.4 mM isopropyl plasmid pETDuet-1-AldTβ-D-1-thiogalactopyranoside were grown in(IPTG) 500 mL at 20 LB ° mediumC for 16– containing18 h. The carbenicillinBL21 (DE3) µ ◦ cells(100 carryingg/ml) the toan expression OD600 of plasmid 0.5 at37 pETDuetC before-1-AldT induction were grown was performed in 500 ml LB with medium 0.4 mM containing IPTG at 37 ◦C for 4 h. All cultures were harvested by centrifugation at 5000× g for 20 min, resuspended in carbenicillin (100 µg/ml) to an OD600 of 0.5 at 37 °C before induction was performed with 0.4 mM IPTG50 mM at 4-(2-hydroxyethyl)-1-piperazineethanesulfonic 37 °C for 4 h. All cultures were harvested acid by (HEPES) centrifugation pH 7 and at lysed 5000 byx g three for passages 20 min, resuspendedthrough a French in 50 pressuremM 4-(2-hydroxyethyl) cell. Soluble fractions-1-piperazineethanesulfonic of the lysates were obtainedacid (HEPES) by centrifugation pH 7 and lysed at × by15,000 three passagesg for 20 min.through His-tagged a French enzymespressure cell. were Soluble purified fractions by subjecting of the lysates the soluble were extractsobtained to by a centrifugation5 mL nickel-nitrilotriacetic at 15000 x g for acid 20 (Ni-NTA) min. His- affinitytagged enzymes column (GE were Healthcare) purified by equilibrated subjecting the with soluble buffer extractscontaining to a 50 5 mMml nickel sodium-nitrilotriacetic phosphate (NaP), acid (Ni 300-NTA) mM NaCl affinity and column 20 mM (GE imidazole. Healthcare) The proteins equilibrated were witheluted buffer isocratically containing with 50 amM final sodium concentration phosphate of 400(NaP), mM 300 imidazole. mM NaCl The and buffer 20 mM of theimidazole. eluates The was proteinsexchanged were to 50eluted mM HEPESisocratically pH 7 with by three a final rounds concentration of centrifugation of 400 mM in Amicon imidazole. Ultra The centrifugal buffer of filters the eluates(Millipore) was withexchanged a 30 kDa to molecular50 mM HEPES weight pH cut-off. 7 by three rounds of centrifugation in Amicon Ultra centrifugalIn order filters to obtain(Millipore) TaManD with withouta 30 kDa a molecular N-terminal weight His-tag, cut- theoff. enzyme obtained after pProEX HTa-Ta0753In orderexpression to obtain TaManD and first without His-tag purificationa N-terminal was His incubated-tag, the enzyme with a TEV obtained Protease after (Promega, pProEX HTa-Ta0753 expression and first His-tag purification was incubated with a TEV Protease (Promega,

Catalysts 2019, 9, 234 10 of 14

Madison, WI, USA). Digestion was performed according to the manufacturer’s protocol with addition of 10% glycerol (v/v, final concentration) and incubation for 1.5 h at 30 ◦C followed by incubation at 4 ◦C overnight. Removal of the His-tag and the TEV Protease was achieved by a second Ni-NTA purification using a 1 mL ZetaSep Ni-NTA affinity column (emp Biotech, Berlin, Germany). The flow-through contained the pure protein and cleavage of the His-tag was verified by SDS-PAGE.

3.2. Protein Analysis Purified proteins were separated by SDS-PAGE using 4–15% Tris-glycine polyacrylamide gels (Bio-Rad Laboratories, Hercules, USA) and visualised with Coomassie Brilliant Blue. For the identification of TaManD, the single protein band at 38 kDa was excised from the gel and analysed by LC ESI MS/MS at the Australian Proteome Analysis Facility (APAF, Macquarie University), as described elsewhere [34]. Protein identification was performed using the mascot software (2.4.1, Matrixscience, Boston, MA, USA). The native molecular mass of the recombinant protein was determined by size exclusion chromatography on an ÄKTA pure FPLC system using a Superdex Increase 200 10/300 GL column (GE Healthcare) equilibrated with 10 mM NaP buffer, 140 mM NaCl, pH 7. A calibration curve was obtained with molecular weight markers thyroglobulin (669 kDa), ferritin (440 kDa), aldolase (158 kDa), conalbumin (75 kDa) and ovalbumin (43 kDa).

3.3. Nuclear Magentic Resonance (NMR) Analysis D-Mannono-1,4-lactone (>95%) was purchased from TCI Chemicals Co. (Tokyo, Japan). For NMR spectra of the sugar acid in different substrate forms, 1 M D-Mannono-1,4-lactone solutions were prepared in 1 M NaOH, 1 M HCl and in ultrapure water. All solutions were incubated for 1 h at room temperature, before each solution was diluted to 100 mM using ultrapure water. Then, 300 µL of each 100 mM solution (equivalent to 5 mg sugar acid) was freeze-dried and resuspended in 500 µL 0.1 M NaP buffer, pH 7. The pH of all three samples was evaluated prior to NMR analysis. The substrate incubated in HCl displayed pH 6, while the substrate in water or NaOH displayed pH 7. After transfer of each sample to 5 mm NMR tubes, 10% D2O and 40 µM 3-(trimethylsilyl)propionic-2,2,3,3-d4 acid (TMSP) chemical shift standard was added. The 1D decoupled 13C NMR spectra of non-isotopically labelled samples were acquired on a 500 MHz Bruker Avance III HD NMR equipped with a BBFO probe at 50 ◦C (323 K), using power-gated proton decoupling with a 90 ◦ pulse with 512 scans and 13 a 3 s relaxation delay between scans. Oxidation of D-[1,6- C2]mannose was performed in reactions containing 5 mM of the substrate, 5 mM NAD+, 20 mM NaP buffer at pH 7 and 13.5 µg puriﬁed AldT. Prior to the addition of puriﬁed enzyme and after 30 min of reaction at 50 ◦C, 1D power-gated proton decoupled 13C NMR spectra were acquired using a 90 ◦ pulse with 4 scans and a 3 s relaxation delay between scans. Visualisation of all spectra was performed with iNMR 6.0 (http://www.inmr.net). After Fourier transformation and automatic phase correction, an exponential visual weighting factor of 1.5 and a smoothing factor of 10 were applied to all spectra.

3.4. Enzyme Activity D-mannono-1,4-lactone was either used after preparation in 50 mM HEPES at pH 7 (indicated as “lactone in buffer”) or after hydrolysis to its free acid form (indicated as “mannonate” or “free sugar acid”). Hydrolysis was performed according to Lamble et al. by preparing 1 M stock solutions of D-mannono-1,4-lactone in 1 M NaOH and incubation at room temperature for 1 h before dilutions were prepared in 50 mM HEPES pH 7 [35]. For temperature optimum, pH optimum and thermostability, activity was quantified using the thiobarbituric acid (TBA) assay, according to a modification by Buchanan et al. [36,37]. Unless stated otherwise, reactions (60 µL) contained 0.5–1.5 µg pure enzyme, ◦ 10 mM mannonate, 50 mM HEPES pH 7 (adjusted at 55 C) and 1 mM CoSO4. Reactions were stopped by incubation on ice or if needed by addition of 6 µL 12.5% trichloracetic acid (TCA). Next, 50 µL of the reaction mixture was oxidised using 125 µL of 25 mM periodic acid in 0.25 M H2SO4 at room temperature for 20 min. The oxidation was stopped by addition of 250 µL of 2% (w/v) sodium arsenite Catalysts 2019, 9, 234 11 of 14 in 0.5 M HCl. Finally, 1 mL of 0.3% TBA in water was added to the samples and the reaction mixture was boiled for 10 min. The formation of 2-keto-3-deoxy sugar acid was determined by reading A549nm using 96-well microtiter plates in a Spectrostar Nano plate reader (BMG Labtech, Ortenberg, Germany) and quantified using the molar extinction coefficient of 67.8 × 103 M−1 × cm−1. The effect of pH on enzyme activity was analysed in duplicate reactions with 1 µg enzyme and 120 mM universal buffer [38]. Reactions were incubated for 2 h at 55 ◦C before they were analysed with the TBA Assay. In reactions with D-mannono-1,4-lactone in buffer, pH decreased during the course of the experiment, which was accounted for in the analysis by measurement of the pH after the reaction. The optimal temperature for enzyme activity was determined in duplicate reactions containing 1 µg enzyme. Duplicate reactions were performed for 1 h at different temperatures before being analysed with the TBA Assay. For thermostability assays, 120 mM universal buffer was used and purified enzyme (1 µg) was incubated in duplicates at temperatures between 55 ◦C and 95 ◦C in the absence of substrate. Samples were removed at different times and the residual activity was measured in reactions with 10 mM mannonate for 45 min at 55 ◦C. The semicarbazide assay was used according to Wichelecki et al. [12] to determine the effect of metal ions, chelators and reducing agents, and the acquisition of Michalis-Menten kinetics (Km, Vmax). Each reaction (60 µL) was incubated with 240 µL semicarbazide reagent at room temperature for 1 h. A250nm was read in a UV transparent microtiter plate (Thermo Scientific, Waltham, MA, USA) using a Spectrostar Nano plate reader. Product formation was quantified using a standard curve of 2-keto-3-deoxygluconate (Sigma-Aldrich, St. Louis, MO, USA) prepared in the same assay buffer. Km, Vmax values were estimated using non-linear fitting in Prism 6 (6.0c, GraphPad software, San Diego, CA, USA). For the effect of metals, chelators and reducing agents, pure enzyme (0.6 µg) was incubated with 1 mM of each additive in 50 mM HEPES pH 7 for 1 h at room temperature. Activity was measured in duplicate reactions containing 10 mM D-mannono-1,4-lactone in 50 mM HEPES pH 7, incubated for 1 h at 55 ◦C and then analysed with the semicarbazide assay. Kinetic data for the determination of Km and Vmax were performed in duplicate reactions (60 µL), containing 1 µg purified TaManD, 50 mM HEPES pH 7, 1 mM CoSO4 and different substrate concentrations (1–50 mM). Linear increase of reaction product was assured over 1 h reaction time. Reactions were performed for 45 min at 55 ◦C before product formation was determined using the semicarbazide assay and the KDG standard curve. Substrate specificity was analysed using 11 different sugar acids (Figure8B). Reactions were performed in duplicate and contained 0.5 µg enzyme, 50 mM HEPES pH 7, 1 mM CoSO4 and 10 mM of each sugar acid. Reactions were incubated for 16 h at 55 ◦C before they were analysed using the semicarbazide assay.

3.5. High Performance Liquid Chromatography (HPLC) Analysis Reactions catalysed by TaManD and standards of mannonate and KDG were analysed using an Agilent 1290 HPLC system connected to a refractive index detector (RID) G1362A (Agilent Technologies, Santa Clara, CA, USA). Samples were analysed on an organic acid column (Agilent HiPlex H+) with ◦ 10 mM H2SO4 as a mobile phase at a ﬂow rate of 0.6 ml/min. The column was heated to 80 C and the RID was set to 55 ◦C. Reactions were stopped by addition of 7 µL 12.5% TCA to a 60 µL reaction. After short centrifugation, 10 µL of the supernatant was used for HPLC analysis.

3.6. Statistical Analysis Statistical analysis was performed for enzyme kinetics and effect of different metals and additives on enzyme activity, by two-tailed unpaired t-tests using Prism 6.

4. Conclusions In this study, we present the ﬁrst puriﬁcation and characterisation of a functional archaeal mannonate dehydratase. The gene encoding for the mannonate dehydratase was found adjacent to a previously described aldohexose dehydrogenase (AldT) gene in the genome of T. acidophilum [15]. Catalysts 2019, 9, 234 12 of 14

Previously, it has been shown that AldT from T. acidophilum selectively oxidises mannose, but the physiological function behind that oxidation was not investigated. Using NMR spectroscopy, we were able to show that mannonate and mannono-1,4-lactone are produced via oxidation of D-mannose by AldT. Kinetic assays conﬁrmed that TaManD was able to convert both the sugar acid and its lactone to the central intermediate KDG at neutral pH, without further help of a lactonase. This resembles the second step of many oxidative pathways studied in archaea, including those for sugars like glucose, galactose, rhamnose, arabinose and xylose [39]. The amino acid sequence of TaManD and those of the putative mannonate dehydratases from F. acidarmanus and F. acidiphilum share high amino acid sequence identity. Although the gene annotations for those mannonate dehydratases indicate a role in the hexuronate metabolism (uxuA), they are all located adjacent to (putative) aldohexose dehydrogenases. It remains to be seen whether a non-phosphorylative pathway starting from mannose, similar to the non-phosphorylative Entner-Doudoroff pathway from glucose and galactose, exists in thermophilic archaea like T. acidophilum, F. acidarmanus and F. acidiphilum.

Supplementary Materials: The following are available online at http://www.mdpi.com/2073-4344/9/3/234/s1, Figure S1: Purification of the Ta0753 gene product (TaManD) after expression in E. coli, Figure S2: TaManD kinetic data Table S1: Overview of genes, potentially encoding for mannonate dehydratases located in the genomic neighbourhood of AldT (Ta0754), Table S2: Oligonucleotides used in this study. Author Contributions: Conceptualization, D.K. and A.S.; methodology, experimental design, D.K., R.W., and A.S.; data collection, D.K.; data analysis, D.K.; writing—original draft preparation, D.K. and A.S.; writing—review and editing, D.K., R.W., and A.S.; supervision, A.S.; project administration, A.S. Funding: D.K. is supported by an international Macquarie University Research Excellence Scholarship (iMQRES). Acknowledgments: We thank the Australian Proteome Analysis Facility (Macquarie University, Sydney, Australia) for their support in performing LC ESI MS/MS experiments and Nicole Cordina from the Macquarie University NMR facility from the Department of Molecular Sciences for her help in performing NMR studies. Conflicts of Interest: The authors declare no conflict of interest.

References

1. Robert-Baudouy, J.; Jimeno-Abendano, J.; Stoeber, F. D-Mannonate and D-altronate dehydratases of Escherichia coli K12. Methods Enzymol. 1982, 90, 288–294. [PubMed] 2. Shulami, S.; Gat, O.; Sonenshein, A.L.; Shoham, Y. The glucuronic acid utilization gene cluster from Bacillus stearothermophilus T-6. J. Bacteriol. 1999, 181, 3695–3704. [PubMed] 3. Mekjian, K.R.; Bryan, E.M.; Beall, B.W.; Moran, C.P. Regulation of hexuronate utilization in Bacillus subtilis. J. Bacteriol. 1999, 181, 426–433. [PubMed] 4. Hugouvieux-Cotte-Pattat, N.; Robert-Baudouy, J. Hexuronate catabolism in Erwinia Chrysanthemi. J. Bacteriol. 1987, 169, 1223–1231. [CrossRef][PubMed] 5. Mandrand-Berthelot, M.-A.; Condemine, G.; Hugouvieux-Cotte-Pattat, N. Catabolism of hexuronides, hexuronates, aldonates, and aldarates. EcoSal Plus 2004, 1, 1–21. [CrossRef][PubMed] 6. Reis, D.; Vian, B.; Roland, J.C. Cellulose-glucuronoxylans and plant cell wall structure. Micron 1994, 25, 171–187. [CrossRef] 7. Lawford, H.G.; Rousseau, J.D. Fermentation of biomass-derived glucuronic acid by pet expressing recombinants of E. coli B. Appl. Biochem. Biotechnol. 1997, 63–65, 221–241. [CrossRef] 8. Peekhaus, N.; Conway, T. What’s for dinner?: Entner-Doudoroff metabolism in Escherichia coli. J. Bacteriol. 1998, 180, 3495–3502. [PubMed] 9. Chang, D.-E.; Smalley, D.J.; Tucker, D.L.; Leatham, M.P.; Norris, W.E.; Stevenson, S.J.; Anderson, A.B.; Grissom, J.E.; Laux, D.C.; Cohen, P.S.; et al. Carbon nutrition of Escherichia coli in the mouse intestine. Proc. Natl. Acad. Sci. USA 2004, 101, 7427–7432. [CrossRef][PubMed] 10. Zhang, Q.; Gao, F.; Peng, H.; Cheng, H.; Liu, Y.; Tang, J.; Thompson, J.; Wei, G.; Zhang, J.; Du, Y.; et al. Crystal structures of Streptococcus suis mannonate dehydratase (ManD) and its complex with substrate: Genetic and biochemical evidence for a catalytic mechanism. J. Bacteriol. 2009, 191, 5832–5837. [CrossRef] [PubMed] Catalysts 2019, 9, 234 13 of 14

11. Qiu, X.; Tao, Y.; Zhu, Y.; Yuan, Y.; Zhang, Y.; Liu, H.; Gao, Y.; Teng, M.; Niu, L. Structural insights into decreased enzymatic activity induced by an insert sequence in mannonate dehydratase from Gram negative bacterium. J. Struct. Biol. 2012, 180, 327–334. [CrossRef][PubMed] 12. Wichelecki, D.J.; Balthazor, B.M.; Chau, A.C.; Vetting, M.W.; Fedorov, A.A.; Fedorov, E.V.; Lukk, T.; Patskovsky, Y.V.; Stead, M.B.; Hillerich, B.S.; et al. Discovery of function in the enolase superfamily: D-mannonate and D-gluconate dehydratases in the D-mannonate dehydratase subgroup. Biochemistry 2014, 53, 2722–2731. [CrossRef][PubMed] 13. Wichelecki, D.J.; Alyxa, J.; Vendiola, F.; Jones, A.M.; Al-obaidi, N.; Almo, S.C.; Gerlt, J.A. Investigating the physiological roles of low-efficiency D-mannonate and D-gluconate dehydratases in the enolase superfamily: pathways for the catabolism of L-gulonate and L-idonate. Biochemistry 2014, 53, 5692–5699. [CrossRef] [PubMed] 14. Rakus, J.F.; Fedorov, A.A.; Fedorov, E.V.; Glasner, M.E.; Vick, J.E.; Babbitt, P.C.; Almo, S.C.; Gerlt, J.A. Evolution of enzymatic activities in the enolase superfamily: D-mannonate dehydratase from Novosphingobium aromaticivorans. Biochemistry 2007, 46, 12896–12908. [CrossRef][PubMed] 15. Nishiya, Y.; Tamura, N.; Tamura, T. Analysis of bacterial glucose dehydrogenase homologs from thermoacidophilic archaeon Thermoplasma acidophilum: finding and characterization of aldohexose dehydrogenase. Biosci. Biotechnol. Biochem. 2014, 68, 2451–2456. [CrossRef][PubMed] 16. Yasutake, Y.; Nishiya, Y.; Tamura, N.; Tamura, T. Structural Insights into unique substrate selectivity of Thermoplasma acidophilum D-aldohexose dehydrogenase. J. Mol. Biol. 2007, 367, 1034–1046. [CrossRef] [PubMed] 17. Kim, S.; Lee, S.B. Catalytic promiscuity in dihydroxy-acid dehydratase from the thermoacidophilic archaeon Sulfolobus solfataricus. J. Biochem. 2006, 139, 591–596. [CrossRef][PubMed] 18. Ashwell, G. [21] Enzymes of glucuronic and galacturonic acid metabolism in bacteria. In Methods in enzymology; Academic Press: Cambridge, MA, USA, 1962; Volume 5, pp. 190–208, ISBN 0076-6879. 19. Portalier, R.; Robert-Baudouy, J.; Stoeber, F. Regulation of Escherichia coli K-12 hexuronate system genes: exu regulon. J. Bacteriol. 1980, 143, 1095–1107. [PubMed] 20. Palleroni, N.J.; Duodoroff, M. Metabolism of Carbohydrate by Pseudomonas Saccharophila III. Oxidation of Arabinose. J. Bacteriol. 1957, 74, 180–185. [PubMed] 21. Watanabe, S.; Saimura, M.; Makino, K. Eukaryotic and bacterial gene clusters related to an alternative pathway of nonphosphorylated L-rhamnose metabolism. J. Biol. Chem. 2008, 283, 20372–20382. [CrossRef] [PubMed] 22. Stephens, C.; Christen, B.; Fuchs, T.; Sundaram, V.; Watanabe, K.; Jenal, U. Genetic analysis of a novel pathway for D-xylose metabolism in Caulobacter crescentus. J. Bacteriol. 2007, 189, 2181–2185. [CrossRef] [PubMed] 23. Wałaszek, Z.; Horton, D. Conformational studies on aldonolactones by NMR spectroscopy. Conformations of d-glucono-, D-mannono, D-gulono-, D-galactono-1,4-lactone in solution. Carbohydr. Res. 1982, 105, 131–143. [CrossRef] 24. Dreyer, J. The role of iron in the activation of mannonic and altronic acid hydratases, two Fe-requiring hydro-lyases. Eur. J. Biochem 1987, 166, 623–630. [CrossRef][PubMed] 25. Robert-Baudouy, J.M.; Jimeno-Abendano, J.; Stoeber, F.R. Individualité des hydrolyases mannonique et altronique chez E. coli K-12. Biochimie 1975, 57, 1–8. [CrossRef] 26. Robert-Baudouy, J.M.; Stoeber, F.R. Purification et propriétés de la D-mannonate hydrolyase d’Escherichia coli. Biochim. Biophys. Acta -Enzymology 1973, 309, 473–485. [CrossRef] 27. Darland, G.; Brock, T.D.; Samsonoff, W.; Conti, S.F. A thermophilic, acidophilic mycoplasma isolated from a coal refuse pile. Science 1970, 170, 1416–1418. [CrossRef][PubMed] 28. Jung, J.H.; Lee, S.B. Identification and characterization of Thermoplasma acidophilum 2-keto-3-deoxy-D-gluconate kinase: A new class of sugar kinases. Biotechnol. Bioprocess Eng. 2005, 10, 535–539. [CrossRef] 29. Kim, S.M.; Paek, K.H.; Lee, S.B. Characterization of NADP+-specific L-rhamnose dehydrogenase from the thermoacidophilic Archaeon Thermoplasma acidophilum. Extremophiles 2012, 16, 447–454. [CrossRef][PubMed] Catalysts 2019, 9, 234 14 of 14

30. Reher, M.; Schönheit, P. Glyceraldehyde dehydrogenases from the thermoacidophilic euryarchaeota Picrophilus torridus and Thermoplasma acidophilum, key enzymes of the non-phosphorylative Entner-Doudoroff pathway, constitute a novel enzyme family within the aldehyde dehydrogenase superfamily. FEBS Lett. 2006, 580, 1198–1204. [PubMed] 31. Yew, W.S.; Fedorov, A.A.; Fedorov, E.V.; Rakus, J.F.; Pierce, R.W.; Almo, S.C.; Gerlt, J.A. Evolution of enzymatic activities in the enolase superfamily: L-fuconate dehydratase from Xanthomonas campestris. Biochemistry 2006, 45, 14582–14597. [CrossRef][PubMed] 32. Carsten, J.M.; Schmidt, A.; Sieber, V. Characterization of recombinantly expressed dihydroxy-acid dehydratase from Sulfobus solfataricus—A key enzyme for the conversion of carbohydrates into chemicals. J. Biotechnol. 2015, 211, 31–41. [CrossRef][PubMed] 33. Sambrook, J.; Russell, D.W.; Russell, D.W. Molecular Cloning: A Laboratory Manual; Cold Spring Harbor Laboratory Press: New York, NY, USA, 2001; Volume 49, ISBN 978-1-936113-42-2. 34. Atack, J.M.; Srikhanta, Y.N.; Fox, K.L.; Jurcisek, J.A.; Brockman, K.L.; Clark, T.A.; Boitano, M.; Power, P.M.; Jen, F.E.C.; McEwan, A.G.; et al. A biphasic epigenetic switch controls immunoevasion, virulence and niche adaptation in non-typeable Haemophilus inﬂuenzae. Nat. Commun. 2015, 6, 1–12. [CrossRef][PubMed] 35. Lamble, H.J.; Milburn, C.C.; Taylor, G.L.; Hough, D.W.; Danson, M.J. Gluconate dehydratase from the promiscuous Entner-Doudoroff pathway in Sulfolobus solfataricus. FEBS Lett. 2004, 576, 133–136. [CrossRef] [PubMed] 36. Buchanan, C.L.; Connaris, H.; Danson, M.J.; Reeve, C.D.; Hough, D.W. An extremely thermostable aldolase from Sulfolobus solfataricus with speciﬁcity for non-phosphorylated substrates. Biochem. J. 1999, 343, 563–570. [CrossRef][PubMed] 37. Skoza, L.; Mohos, S. Stable thiobarbituric acid chromophore with dimethyl sulphoxide. Application to sialic acid assay in analytical de-O-acetylation. Biochem. J. 1976, 159, 457–462. [CrossRef][PubMed] 38. Britton, H.T.S.; Robinson, R.A. CXCVIII.—Universal buffer solutions and the dissociation constant of veronal. J. Chem. Soc. 1931, 1456–1462. [CrossRef] 39. Bräsen, C.; Esser, D.; Rauch, B.; Siebers, B. Carbohydrate metabolism in archaea: current insights into unusual enzymes and pathways and their regulation. Microbiol. Mol. Biol. Rev. 2014, 78, 89–175. [CrossRef][PubMed]

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