Journal of Bacteriology, September 2004, p . 6198-6207, Vol . 186, No . 18

Novel Xylose Dehydrogenase in the Halophilic Archaeon Haloarcula marismortui

Ulrike Johnsen and Peter Schönheit^*

Institut für Allgemeine Mikrobiologie, Christian-Albrechts-Universität Kiel, Kiel, Germany

Received 3 March 2004/ Accepted 21 June 2004

 
During growth of the halophilic archaeon Haloarcula marismortui on D-xylose, a specific D-xylose dehydrogenase was induced.The enzyme was purified to homogeneity . It constitutes a homotetramerof about 175 kDa and catalyzed the oxidation of xylose withboth NADP⁺ and NAD⁺ as cosubstrates with 10-fold higher affinityfor NADP⁺ . In addition to D-xylose, D-ribose was oxidized atsimilar kinetic constants, whereas D-glucose was used with about70-fold lower catalytic efficiency [k_cat/K_m] . With the N-terminalamino acid sequence of the subunit, an open reading frame [ORF]—codingfor a 39.9-kDA protein—was identified in the partiallysequenced genome of H . marismortui . The function of the ORFas the gene designated xdh and coding for xylose dehydrogenasewas proven by its functional overexpression in Escherichia coli.The recombinant enzyme was reactivated from inclusion bodiesfollowing solubilization in urea and refolding in the presenceof salts, reduced and oxidized glutathione, and substrates.Xylose dehydrogenase showed the highest sequence similarityto glucose-fructose oxidoreductase from Zymomonas mobilis andother putative bacterial and archaeal oxidoreductases . Activitiesof xylose isomerase and xylulose kinase, the initial reactionsof xylose catabolism of most bacteria, could not be detectedin xylose-grown cells of H . marismortui, and the genes thatencode them, xylA and xylB, were not found in the genome ofH . marismortui . Thus, we propose that this first characterizedarchaeal xylose dehydrogenase catalyzes the initial step inxylose degradation by H . marismortui.

 
The utilization of sugars, in particular of hexoses and hexose polymers and—to a lesser extent—of pentoses, hasbeen reported for various species in the domain Archaea . Sofar, only the catabolic pathways of hexoses and glucose polymers[e.g., maltose and starch] have been studied in detail in particularin hyperthermophilic, thermoacidophilic, and extremely halophilic archaea . Comparative analyses of glucose degradation pathwaysin these organisms revealed that the classical Embden-Meyerhof-[EM] or Entner-Doudoroff- [ED] pathway found in bacteria isnot operative in archaea; they use instead modified versionsof these pathways as follows [for reviews, see references 31and 41] . In hyperthermophilic eury- and crenarchaeota, glucose degradation proceeds predominantly via modified EM pathways, which differ from the classical EM pathway by the presence ofseveral unusual glucokinases [ADP or ATP dependent] and 6-phosphofructokinases [ADP, ATP, or PP_i dependent], novel enzymes of glucose-6-phosphateisomerization and of glyceraldehyde-3-phosphate oxidation, andpyruvate kinases with reduced regulatory potential [15, 18,41].

In thermoacidophilic archaea, Sulfolobus and Thermoplasma spp., glucose is degraded via a nonphosphorylated version of the ED pathway [22, 31, 41] by which glucose is oxidized to glyceratevia the nonphosphorylated intermediates gluconate and 2-keto-3-deoxygluconate[KDG] involving glucose dehydrogenase, gluconate dehydratase,and KDG aldolase . Glycerate is then phosphorylated via a specifickinase to 2-phosphoglycerate, which is further converted topyruvate via enolase and pyruvate kinase . In halophilic archaea,e.g., Halococcus, Haloarcula, and Haloferax spp., a modified, semiphosphorylated ED pathway is operative in which—asin thermoacidophiles—glucose is converted to KDG . However,KDG is then phosphorylated to 2-keto-3-deoxy-6-phosphogluconateby KDGkinase . Further degradation of 2-keto-3-deoxy-6-phosphogluconateproceeds via reactions of the conventional phosphorylated EDpathway found in bacteria [19, 45].

In contrast to hexose metabolism, the catabolic pathways of pentoses have not been studied in detail in the domain Archaea. The utilization of pentoses, e.g., xylose, ribose, and arabinose, has been reported for several halophiles, e.g., Halococcus, Haloarcula, and Halobacterium spp., and for Sulfolobus species[30, 40], rather than for the majority of hyperthermophiles.No studies of growth on pentoses or analyses of the enzymesinvolved in pentose degradation by these organisms have beenreported.

In the domain Bacteria, the pathways for the degradation of pentoses, in particular, D-xylose, have been studied in detailin many species, including Escherichia coli, Salmonella entericaserovar Typhimurium, Lactobacillus pentosus, Lactococcus lactis,Bacillus spp., Staphylococcus xylosus, Bacteroides xylanolyticus,and Tetragenococcus halophilus . Degradation of xylose by these organisms, e.g., by E . coli, starts with its uptake via specific high- or low-affinity transport systems . Via xylose isomerase, xylose is then isomerized to xylulose, which is phosphorylatedto xylulose-5-phosphate by the activity of xylulose kinase.The genes encoding xylose transporters, xylose isomerase [xylAgene], and xylulose kinase [xylB gene], which are arranged inan operon, are induced by xylose mediated by the transcriptional regulator XylR . Further degradation of xylulose-5-phosphate, proceeds—depending on the organism—either via thepentose phosphate cycle, the phosphoketolase pathway, or—asin Bacteroides spp . [4]—via a combination of both pentosephosphate and the EM pathway . Thus, the most common initialreactions of bacterial xylose catabolism involve xylose isomeraseand xylulose kinase [4, 13, 23, 29, 36].

Xylose isomerase and the gene that encodes it, xylA, have been characterized in many bacteria, including the hyperthermophile Thermotoga maritima, as well as in eucarya . The enzymes from both domains are of significant industrial interest since theyalso catalyze, as a side activity, the isomerization of glucoseto fructose, a reaction that constitutes the last step in the large-scale industrial process of the production of sweetenersfrom starch [3].

So far, activities of xylose isomerase and xylulose kinase have not been reported in any species of the archaeal domain . Further, homologs to the bacterial xylA and xylB genes could not be found in archaeal genomes . Thus, it might be speculated that the initial steps in xylose degradation by archaea are different from the common mechanism found in most bacteria.

In this communication, we report on studies of the growth ofthe halophilic archaeon Haloarcula marismortui on xylose . Evidence is presented that the first step in xylose degradation by this organism is oxidation of xylose to xylonate via a xylose-inducible NADP⁺-reducing D-xylose dehydrogenase . This first archaeal xylosedehydrogenase was purified, and the gene that encodes it, xdh,was identified in available sequenced contigs of H . marismortui.This enzyme represents a novel type of xylose dehydrogenase,showing high similarity to glucose-fructose oxidoreductase [GFOR]from Zymomonas mobilis.

 
Growth of H . marismortui and preparation of cell extracts. H . marismortui [DSM 3752] [24] was obtained from the DeutscheSammlung von Mikroorganismen und Zellkulturen [Braunschweig, Germany] . The organism was grown aerobically at 37°C in500-ml Erlenmeyer flasks filled with 50 ml of medium containing25 mM xylose, 0.05% yeast extract, 250 g of NaCl per liter,20 g of MgSO₄ · 7H₂O per liter, 19.5 g of morpholineethanesulfonicacid [MES] per liter, 2 g of KCl per liter, 1 g of Na-glutamateper liter, and 3 g of Na-citrate per liter; 10 ml of the vitaminsolution described by Staley [35]; and 10 ml of a trace element solution containing [per liter] 1.5 g of EDTA, 0.01 g of Na₂MoO₄ · 2H₂O, 0.5 g of MnSO₄ · H₂O, 0.1 g of FeSO₄ ·7H₂O, 0.1 g of CoCl₂, 0.1 g of ZnSO₄, and 0.01 g of CuSO₄ ·5H₂O . The pH was adjusted to 7.35 with 10 N NaOH . Growth wasfollowed by measuring optical density at 578 nm [OD₅₇₈] . Duringgrowth, samples were removed and centrifuged and the supernatantswere analyzed for xylose and xylonate, as indicated . Extractswere prepared from late-log-phase cells by sonication as describedpreviously [19], and enzyme activities were determined as describedin the section on enzyme assays . Protein was determined by thebiuret method with bovine serum albumin as the standard [5].

Induction of xylose dehydrogenase in H . marismortui was followed in 2,000-ml Erlenmeyer flasks filled with 400 ml of medium containing xylose [25 mM], yeast extract [2.5 g/liter], and Casamino Acids [5 g/liter]; cells previously grown on glucose [see reference 19] were used as an inoculum [10%] . At the times indicated, 60- to 80-ml samples were removed and centrifuged [2,600 x g,10 min, 4°C] and the cell pellets were suspended in 1 mlof 0.1 M Tris-HCl, pH 7.5, containing 250 g of NaCl/liter . Cellextracts were prepared by sonication [19], followed by centrifugationat 12,000 x g for 10 min . The supernatants were analyzed forxylose dehydrogenase activities . The protein concentration ofcell extracts was determined by the biuret method.

Purification of xylose dehydrogenase from H . marismortui. Xylose dehydrogenase was purified from H . marismortui after growth of the organism in a medium [see above] containing 25mM xylose and 0.1% yeast extract in a 10-liter Biostat fermentor. Extract was prepared from 4 g of cells, which were suspendedin 100 mM Tris-HCl, pH 8.8, containing 2 M [NH₄]₂SO₄ and 20mM MgCl₂ [buffer A] . Cells were disrupted by passage througha French pressure cell at 1.3 x 10⁸ Pa . Cell debris and unbrokencells were removed by centrifugation for 90 min at 100,000 xg at 4°C . The 100,000 x g supernatant was applied to a SepharoseCL 4B column [1.6 by 60 cm] that had been equilibrated with buffer A . Protein was eluted with a decreasing [NH₄]₂SO₄ gradientfrom 2 to 0 M in buffer A . Fractions containing the highest xylose dehydrogenase activity [1.6 to 1.4 M [NH₄]₂SO₄] werepooled, adjusted to 2 M [NH₄]₂SO₄, and applied to a Phenyl Sepharosecolumn [2.6 by 10 cm] equilibrated with buffer B [50 mM Tris-HCl,pH 8.5, containing 2 M [NH₄]₂SO₄ and 20 mM MgCl₂] . Protein waseluted with a linear gradient of buffer B to 50 mM Tris-HCl,pH 8.5, containing 20 mM MgCl₂ and 10% glycerol . Fractions containingthe highest xylose dehydrogenase activity [1.04 to 0.95 M [NH₄]₂SO₄] were pooled and concentrated to 600 µl by ultrafiltration [cutoff, 20 kDa] . The concentrated protein solution was appliedto a Superdex 200 HiLoad gel filtration column [1.6 by 60 cm]that had been equilibrated with 50 mM Tris-HCl, pH 8.5, containing20 mM MgCl₂, 10% glycerol, and 100 mM NaCl . In this buffer,the enzyme was stable in the absence of a high salt [KCl orNaCl] concentration . Eluted fractions containing xylose dehydrogenaseactivity indicated essentially pure protein and were storedat –20°C . The purity of the preparations was checkedby sodium dodecyl sulfate-polyacrylamide gel electrophoresis[SDS-PAGE] in 12% gels in accordance with standard procedures[21] . During the purification procedure, protein concentrationswere determined by the Bradford method with bovine serum albuminas the standard [7].

Cloning and expression of xylose dehydrogenase from H . marismortui in E . coli. On the basis of the N-terminal amino acid sequence, an openreading frame [ORF] was identified by a BLASTP search in contig97 of the partially sequenced genome of H . marismortui [P . Zhang,W . V . Ng, and S . DasSarma, personal communication, 2003] . TheORF was characterized as the xdh gene, encoding xylose dehydrogenase,by its functional overexpression in E . coli . The gene was amplifiedfrom genomic DNA of H . marismortui by PCR and cloned into pET17b[Novagen] via two restriction sites [NdeI and BamHI] createdwith the primers 5'-GACGACAGTCATATGAACGTTG-3' and 5'-CAAAAAATCTGGATCCGGTTTC-3'[restriction sites are underlined] . The vector pET17b-xdh wastransformed into E . coli BL21 codon plus[DE3]-RIL [Stratagene].For expression, cells were grown in Luria-Bertani medium at37°C . Expression was initiated by the addition of isopropyl-ß-D-thiogalactopyranoside [IPTG; final concentration, 0.4 mM] . After 18 h of further growth, cells were harvested by centrifugation.

Solubilization, refolding, and purification of recombinant xylose dehydrogenase. Recombinant xylose dehydrogenase, which was expressed in inclusionbodies, was solubilized and refolded as described by Connariset al . [11], with modifications . The E . coli cell pellets weresuspended in 20 mM Tris-HCl, pH 7.5, containing 2 mM EDTA, 3M KCl, and 10% glycerol [buffer C] . The cell suspension wastreated with 100 µg of lysozyme per ml and 0.1% [vol/vol]Triton X-100 and incubated at 30°C for 60 min, followed by incubation on ice for 15 min . The suspension was then sonicated and centrifuged at 40,000 x g for 30 min at 4°C . The insolublefraction was washed twice in buffer C, yielding inclusion bodiesand insoluble cell fragments . The insoluble fraction was dissolvedin 20 mM Tris-HCl, pH 7.5, containing 2 mM EDTA, 8 M urea, and50 mM dithioerythritol . Solubilization was carried out at 37°Cfor 1 h . Refolding was initiated by slowly diluting the suspensionin 20 mM Tris-HCl, pH 7.5, containing 3 M KCl, 2 mM EDTA, 10%glycerol, 2 mM xylose, 0.1 mM NADP⁺, 3 mM reduced glutathione,and 0.3 mM oxidized glutathione to a final concentration ofabout 30 µg of protein per ml . After incubation for 6days at 4°C, the renatured protein was concentrated 250-foldby ultrafiltration [cutoff, 30 kDa] . The concentrated proteinsolution was applied to a Superdex 200 HiLoad 16/60 gel filtrationcolumn that had been equilibrated with 50 mM Tris-HCl, pH 8.5,containing 20 mM MgCl₂, 10% glycerol, and 100 mM NaCl . Elutedfractions containing xylose dehydrogenase activity were pooledand applied to a Phenyl Resource column [1 ml], equilibratedwith buffer B . Protein was eluted with a linear [NH₄]₂SO₄ gradientto 0 M in 50 mM Tris-HCl, pH 8.5, containing 20 mM MgCl₂ and10% glycerol . Essentially pure enzyme was eluted at about 1 M [NH₄]₂SO₄ . The purity of the preparations was checked by SDS-PAGE,and protein concentrations were determined by the Bradford method[7].

Enzyme assays. All enzyme assays were done at 37°C . One unit of enzymeactivity corresponds to the conversion of 1 µmol of substrateconsumed or product formed per min.

Xylose dehydrogenase activity [xylose + NADP⁺ xylonate + NADPH+ H⁺] was assayed by measuring the rate of reduction of NADP⁺ at 365 nm . The standard assay mixture contained 100 mM Tris-HCl[pH 8.3], 1.5 M KCl, 1 mM NADP⁺, 10 mM xylose, and protein.

Glucose dehydrogenase activity was tested as glucose-dependent reduction of NADP⁺ . The assay mixture contained 100 mM Tris-HCl [pH 8.3], 1.5 M KCl, 10 mM glucose, 1 mM NADP⁺, and protein.

Xylose isomerase activity was tested as xylose-dependent formation of xylulose . The assay mixture contained 100 mM Tris-HCl [pH8.5], 1 M KCl, 1 mM CoCl₂, 5 mM MnSO₄, 5 mM MgCl₂, 10 to 100mM xylose or glucose, and protein . During incubation [0 to 20 min], aliquots were taken and the reaction was stopped by addition of trichloroacetic acid to a final concentration of 10% . After centrifugation, xylulose was quantified by the cysteine-carbazole method [17] . Crude extract from xylose-grown E . coli cells servedas a positive control [12].

Xylulose-5-phosphate kinase activity was tested as the ATP-dependent decrease in xylulose . The assay mixture contained 100 mM Tris-HCl [pH 8.5], 1 M KCl, 10 mM MgCl₂, 4 mM cysteine-HCl, 10 mM ATP, 6 mM xylulose, and protein . During incubation [0 to 30 min], aliquots were taken and the reaction was stopped by additionof trichloroacetic acid to a final concentration of 10% . After centrifugation, xylulose was quantified by the cysteine-carbazole method . Crude extract from xylose-grown E . coli cells served as a positive control.

Temperature and pH dependence, salt effects, and cation specificity. The temperature dependence of xylose dehydrogenase was measured between 20 and 60°C in 50 mM Tris-HCl, pH 8.3, containing1.5 M KCl, 1 mM NADP⁺, 10 mM xylose, and protein . The pH dependence of the enzyme was measured between pHs 4.4 and 10.3 at 37°Cwith either piperazine [pHs 4.9 to 6.0], bis-Tris [pHs 6.0 to7.5], Tris-HCl [pHs 7.5 to 9.3], or piperazine [pHs 9.3 to 10.8],each at 20 mM, containing 1.5 M KCl, 1 mM NADP⁺, 10 mM xylose,and protein . The effects of salts [0 to 200 mM MgCl₂, 0 to 3.5 M KCl, and 0 to 3.5 M NaCl] on xylose dehydrogenase activitywere tested at 37°C in 20 mM Tris-HCl, pH 8.3, containing1 mM NADP⁺, 10 mM xylose, and protein.

Substrate specificity. The substrate specificity of xylose dehydrogenase was testedat 37°C in 20 mM Tris-HCl, pH 8.3, containing 1.5 M KClwith D isomers of the sugars in the presence of NADP⁺ at 10mM each xylose and ribose; 1 mM NADP⁺; 100 mM glucose, 1 mMNADP⁺; 100 mM each galactose, fructose, and arabinose; and 2mM NADP⁺ . For the determination of apparent K_m and V_max valuesfor sugars and the cosubstrates NADP⁺ and NAD⁺, the followingconcentrations were used: xylose or ribose, 0 to 10 mM with1 mM NADP⁺; NADP⁺, 0 to 1 mM with 10 mM xylose or ribose; NAD⁺,0 to 3 mM with 10 mM xylose or ribose; glucose, 0 to 100 mMwith 1 mM NADP⁺; NADP⁺, 0 to 1 mM with 100 mM glucose.

Analytical assays. Gel filtration chromatography was carried out with a flow rateof 1 ml/min on a Superdex 200 HiLoad column [1.6 by 60 cm].The column was equilibrated with 50 mM Tris-HCl, pH 8.5, containing20 mM MgCl₂, 10% glycerol, and 100 mM NaCl . HWM and LWM gelfiltration calibration kits [Amersham Biosciences, Amersham,England] were used as the standards.

The concentration of xylose was determined by using the orcinol assay [8] . The extinction coefficient [] at 546 nm was 4,800M^–1 cm^–1 . The concentration of xylonate was determinedby high-performance liquid chromatography with an Aminex HPX87Hcolumn [Bio-Rad, Richmond, Calif.] operating at 37°C . Sampleswere diluted 1:5 in 5 mM H₂SO₄, boiled for 30 min, centrifuged,passed through a 0.2-µm-pore-size filter, and loaded ontothe column . Xylonate was eluted with 5 mM H₂SO₄ at a flow rateof 0.6 ml/min and then monitored with a differential refractometerat 210 nm.

 
Growth of H . marismortui on xylose and induction of xylose dehydrogenase. H . marismortui was grown on a medium containing 25 mM D-xyloseand 0.05% yeast extract, with xylose-grown cells [10%] as theinoculum . The cells grew exponentially with a doubling timeof about 20 h up to a OD₅₇₈ of about 1 . During growth, xylosewas consumed and small amounts of xylonate were formed [Fig.1] . In the absence of xylose, the cells grew with a doublingtime of about 30 h up to a OD₅₇₈ of about 0.5 because of theyeast extract present in the medium [data not shown].

 
To identify the first enzymes of xylose degradation, extractsof xylose-grown H . marismortui cells were analyzed for xylose isomerase and xylulose kinase, the initial enzymes of xylose degradation by bacteria . Neither of these activities could be detected . As a control, both enzyme activities [xylose isomerase,40 mU/mg; xylulose kinase, 420 mU/mg] were found in xylose-grownE . coli cells under conditions identical to those used for H. marismortui.

Since during growth on xylose small amounts of xylonate were formed, we looked for enzymes catalyzing the dehydrogenationof xylose . Indeed, extracts of xylose-grown cells catalyzedthe NADP⁺-dependent conversion of xylose to xylonate at a specificactivity of 0.15 U/mg with an apparent K_m for xylose of 0.95mM . Extracts of xylose-grown cells also catalyzed the oxidationof glucose with NADP⁺ to gluconate, however, at a 70-fold lowercatalytic efficiency [apparent V_max, 0.03 U/mg; K_m, 15 mM],indicating that xylose-grown H . marismortui cells contain aspecific xylose dehydrogenase different from glucose dehydrogenase. Glucose-grown cells of H . marismortui also contained xylose dehydrogenase activity, however, with about 10-fold lower catalytic efficiency [apparent V_max, 0.03 U/mg; K_m, 2 mM] compared tothat of xylose-grown cells, suggesting that xylose dehydrogenasewas induced during growth on xylose.

Induction of xylose dehydrogenase was demonstrated during growth of H . marismortui on xylose with cells pregrown on glucose as an inoculum . Because of the higher concentrations of yeast extracts and Casamino Acids present in the medium, the cells grew witha shorter doubling time [about 12 h] to significantly highercell densities and the amount of xylose consumed increased [Fig. 2] . During growth, the NADP⁺-dependent xylose dehydrogenaseactivity increased up to fivefold parallel to xylose consumption,indicating that the enzyme is induced by xylose and probablyrepresents the first reaction of xylose catabolism in H . marismortui.

 
Purification of xylose dehydrogenase from H . marismortui. Xylose dehydrogenase was purified from cell extract of xylose-grown H . marismortui cells by only three chromatographic steps . The most efficient purification step was hydrophobic interaction chromatography on Phenyl Sepharose, resulting in 130-fold enrichment. With the entire procedure, the enzyme was purified about 210-fold,to a specific activity of 100 U/mg with a yield of 10% [Table 1] . The purified protein was electrophoretically homogeneous as judged by denaturing SDS-PAGE [Fig . 3] . Thus, xylose dehydrogenaserepresents about 0.5% of the cellular protein of H . marismortui.

 
Molecular and catalytic properties. The apparent molecular mass of native xylose dehydrogenase determinedby gel filtration on Superdex 200 was 175 ± 15 kDa . SDS-PAGErevealed only one subunit with an apparent molecular mass of57 ± 3 kDa [Fig . 3] . This value is significantly overestimated,as has been observed for various halophilic enzymes [see Discussion].Recombinant xylose dehydrogenase [see below], showed an apparentmolecular mass on SDS-PAGE of 57 ± 3 kDa, although thecalculated molecular mass is 39.9 kDa . We propose that nativexylose dehydrogenase is a homotetrameric [₄] enzyme.

The purified enzyme catalyzed the oxidation of D-xylose withboth NADP⁺ and NAD⁺ . The rate dependence of the enzyme on xylose,NADP⁺, and NAD⁺ followed Michaelis-Menten kinetics with apparentK_m values of 1.2, 0.15, and 0.9 mM, respectively . The correspondingV_max values were about 100, 92, and 80 U/mg . The sixfold higherapparent K_m for NAD⁺ compared to NADP⁺ indicates that NADP⁺ is the preferred electron acceptor . In addition to D-xylose,various other pentoses and hexoses [all D isomers] were testedas substrates for the dehydrogenase with NADP⁺ as the electronacceptor . The apparent K_m values, V_max, k_cat values, and catalyticefficiencies [k_cat/K_m] are given in Table 2 . The highest catalytic efficiency was obtained with xylose and NADP⁺ . D-Ribose wasalso accepted by the enzyme at high efficiency, whereas D-arabinosewas not oxidized at significant rates, indicating that the configurationchange at C-2 significantly affects enzyme activity . D-Glucosewas oxidized with a catalytic efficiency 70-fold lower thanthat with which xylose was oxidized, thus defining the enzymeas a specific xylose dehydrogenase [Fig . 4 A and B] . Galactose oxidation was less efficient than glucose oxidation, and almostno activity was found with fructose [Table 2].

 
Effects of salt, pH, and temperature on xylose dehydrogenase activity. The activity of xylose dehydrogenase from H . marismortui was strongly stimulated by high concentrations of NaCl or KCl andby moderate concentrations of MgCl₂ . Maximal activities were obtained at about 1.5 M both KCl and NaCl and at 100 mM MgCl₂ [Fig . 5] . The rate dependence of the enzyme on pH and temperaturewas tested in the presence of 1.5 M KCl . The pH optimum wasabout pH 8.3, and 50% activity was found at pHs 7 and 9 . Xylose dehydrogenase activity increased exponentially with temperature between 25 and 45°C; from the corresponding linear partof an Arrhenius plot, an activation energy of 64 kJ/mol wascalculated . The highest catalytic activity of xylose dehydrogenasewas found at 50°C.

 
Identification, sequence analysis, and cloning of the gene encoding xylose dehydrogenase from H . marismortui and its functional overexpression in E . coli. On the basis of the N-terminal amino acid sequence determinedfrom the subunit of xylose dehydrogenase, MNVDALTGGFDRRDWQEQTATDNPVRFAA,an ORF that exactly matches the 29 N-terminal amino acid residueswas identified by BLASTP search in contig 97 of the partiallysequenced genome of H . marismortui [Zhang et al., personal communication].The ORF contains 1,083 bp coding for a polypeptide of 360 aminoacids with a calculated molecular mass of 39.9 kDa; the proteincontains large amounts of negatively charged amino acids, 10%Asp and 11% Glu, which is typical for halophilic enzymes [10,27], and had a predicted pI of 4.2 . The G+C content of the ORFis 62 mol% . The coding sequence starts with ATG and stops withTGA . A putative archaeal box A [TATA box] promoter signal [5'-TAATAT-3']was identified between positions –23 and –28 upstreamfrom the ATG start codon [25, 37] . Immediately upstream of theinitiation codon, a putative ribosome binding site with the sequence 5'-GTGGT-3' is present [32] . Downstream of the gene,a pyrimidine-rich sequence beginning at position 1069 and a short inverted repeat located between positions 1112 and 1121 were identified, indicating a transcription termination site[38].

The ORF was characterized as the xdh gene encoding xylose dehydrogenaseby its functional overexpression in E . coli . The xdh gene wasamplified by PCR and cloned into the vector pET17b . The recombinantplasmid was used to transform E . coli BL21 codon plus[DE3]-RIL.After induction with IPTG, a polypeptide of about 59 kDa wasoverexpressed, which was recovered almost completely in inclusionbodies . The protein purified from inclusion bodies was a catalyticallyactive, extremely halophilic xylose dehydrogenase.

Solubilization, refolding, and purification of recombinant xylose dehydrogenase. Recombinant xylose dehydrogenase was purified from inclusionbodies by dissolving with 8 M urea in the presence of dithioerythritol,followed by refolding with a buffer containing a high salt concentration[3 M KCl], substrates, and glutathione [see Materials and Methods].Maximal xylose dehydrogenase activity was obtained after 6 daysof incubation . The refolded activated xylose dehydrogenase waspurified by chromatography on Superdex and Phenyl Resource.The recombinant xylose dehydrogenase showed molecular and kineticproperties almost identical to those of the enzyme purified from H . marismortui [Table 2] . The molecular masses of the nativeenzyme and subunits were 180 ± 10 and 59 ± 3 kDa [Fig . 3], respectively, and the apparent K_m values were verysimilar for xylose, ribose, glucose, and NADP⁺ [data not shown];however, the apparent V_max of the recombinant enzyme was significantly[about 40%] lower.

 
In the present communication, we describe the purification and characterization of the first archaeal D-xylose dehydrogenaseand the gene that encodes it from the halophilic archaeon H.marismortui . The enzyme was induced during growth on xylose,suggesting that xylose dehydrogenase represents the initialenzyme of the xylose degradation pathway in this archaeon . Theenzyme constitutes a novel type of xylose dehydrogenase related to GFOR from Z . mobilis.

Molecular and kinetic properties. Xylose dehydrogenase was characterized as a homotetrameric enzymeof about 175 kDa; the calculated subunit molecular mass is 39.9kDa . The apparent molecular mass of subunits on SDS-PAGE ofabout 57 kDa obtained for xylose dehydrogenase was overestimatedas reported for several halophilic proteins, probably becauseof the presence of large amounts of negatively charged aminoacids . The same degree of overestimation as described for xylosedehydrogenase was observed with glucose dehydrogenase from Haloferaxmediterranei . The enzyme has a calculated molecular mass of39.3 kDa [27] and showed an apparent molecular mass on SDS-PAGEof 53 ± 3 kDa [6].

Xylose dehydrogenase showed dual cofactor specificity for pyridine nucleotides with a high preference for NADP⁺ over NAD⁺, indicatingthat NADP⁺ is the physiological electron acceptor . The enzymecatalyzed the oxidation of xylose, ribose, and glucose; however,the catalytic activity for the latter was about 70-fold lower.The archaeal xylose dehydrogenase can be discriminated fromarchaeal glucose dehydrogenases characterized from various organismsincluding Haloferax, Sulfolobus, Thermoplasma, and Thermoproteusspp., which all show xylose dehydrogenase activity . These archaealglucose dehydrogenases are tetrameric or dimeric enzymes composedof 40-kDa subunits, show dual cofactor specificity for NADP⁺ and NAD⁺ with a high preference for NADP⁺, and utilize variousaldoses [hexose and pentoses] including xylose in addition toglucose . However, in contrast to xylose dehydrogenase from H.marismortui, all archaeal glucose dehydrogenases showed significantlyhigher catalytic efficiencies for NADP⁺-dependent oxidationof glucose compared to that of xylose [6, 22, 33, 34].

Few reports of purified xylose dehydrogenases from Eucarya and Bacteria are available . An NADP⁺-specific xylose dehydrogenasefrom pig liver was characterized [46] . The enzyme is a homodimercomposed of 32-kDA subunits showing the highest catalytic activitywith xylose but also accepts ribose and glucose at about 7-or 30-fold lower catalytic efficiency . The enzyme was specificfor NADP⁺ and did not reduce NAD⁺ . Recently, Aoki et al . [1]demonstrated that the NADP⁺-dependent D-xylose dehydrogenase of pig liver is identical to dimeric dihydrodiol dehydrogenase [DD] . Copurification of DD activity and xylose dehydrogenaseactivity from pig liver, molecular mass and kinetic analyses,and inhibitor studies showed that the two enzymes are identical[1] . Dimeric DDs catalyze the NADP⁺-dependent oxidation of various aromatic hydrocarbons, e.g., naphthalene dihydrodiol, to the corresponding catechols and also the oxidation of various sugars, with xylose as the most effective sugar substrate . The oxidationrate of naphthalene dihydrodiol was about twofold higher thanthat of xylose [1], suggesting that dihydrodiols are the preferred substrates of the dimeric DD-xylose dehydrogenase . The genes coding for various mammalian dimeric DD-xylose dehydrogenases, including pig liver, rabbit lens, human intestine, and monkeykidney, were sequenced [2], and thus, sequences of eucaryal xylose dehydrogenases are known.

In bacteria, oxidation of xylose to xylonate has been reportedfor several species [9]; an NAD⁺-reducing xylose dehydrogenaseactivity was reported for two Caulobacter species [28]; however—toour knowledge—the only xylose dehydrogenase from bacteriacharacterized to some detail is the enzyme from Arthrobactersp . The enzyme was induced by xylose and showed a high specificityfor xylose [apparent K_m, 17.4 mM] and for NAD⁺ as an electronacceptor . Other pentoses and hexoses, as well as NADP⁺ as acofactor, were not accepted as substrates [44] . The amino acid sequence of the enzyme has not been reported.

Thus, both characterized eucaryal and bacterial D-xylose dehydrogenasesshowed significant differences in molecular and kinetic propertiescompared to the archaeal xylose dehydrogenase from H . marismortui.

Sequence comparison and phylogenetic analysis of archaeal xylose dehydrogenase. On the basis of the N-terminal amino acid sequence of the subunit,the xdh gene encoding the xylose dehydrogenase from H . marismortuiwas identified in contig 97 of the partially sequenced genomeof the organism by functional overexpression in E . coli . BLASTPsearches of nonredundant databases with the deduced amino acidsequence of the xdh gene from H . marismortui revealed varioushits . Almost all of them were putative oxidoreductases or dehydrogenases.The highest degrees of similarity were found with bacterialGFOR from Z . mobilis [41%] and hypothetical GFORs from the bacteriaDeinococcus radiodurans, Caulobacter crescentus, Streptococcus pneumoniae, and Bacillus halodurans [35 to 40%] . Similarities of 31 to 34% with the putative archaeal oxidoreductases or dehydrogenases from Pyrococcus sp., Sulfolobus solfataricus, and Thermoplasmasp . were found, the best scores being obtained for the putativedehydrogenases from Pyrococcus furiosus [PF1919] and S . solfataricus[SSO3015] . Lower degrees of similarity were found with characterizedeucaryal xylose dehydrogenases-dimeric DDs [28 to 29%] and witharchaeal glucose dehydrogenases [17 to 21%] from halophiles[Haloferax and Halobacterium spp.] and thermoacidophiles [Sulfolobusand Thermoplasma spp.] . For accession numbers, see the legendto Fig . 7.

 
The most similar enzyme of xylose dehydrogenase from H . marismortui, i.e., GFOR from Z . mobilis, is a homotetrameric enzyme composed of 40-kDa subunits and containing tightly bound cofactor NADP⁺. The enzyme catalyzes the coupled intermolecular oxidation-reduction of glucose and fructose to form gluconolactone and sorbitol. The periplasmic enzyme is synthesized as a precursor with an N-terminal signal peptide of 52 amino acid residues [14] . A multiple-sequence alignment of xylose dehydrogenase of H . marismortui, mature GFOR from Z . mobilis, and other putative bacterial oxidoreductasesis given in Fig . 6 . The alignment includes a prediction of secondarystructure, which is in accordance with the crystal structureof GFOR from Zymomonas oxidoreductase [20] . Sequence comparisonsindicate a variety of conserved regions including a typicalßß dinucleotide binding pocket [Rossmanfold, amino acids 26 to 57] [16, 43] and—with few deviations—therecently postulated consensus sequence for a novel class ofdehydrogenases, including the highly conserved EKP motif [42] [amino acids 113 to 122] . Although xylose dehydrogenase showsa high degree of similarity to GFOR from Z . mobilis, the twoenzymes catalyze different reactions; xylose dehydrogenase isa dehydrogenase with dual cofactor specificity for NADP⁺ andNAD⁺, whereas GFOR catalyzes the coupled oxidation-reductionof glucose and fructose with tightly bound NADP⁺, i.e., in theabsence of added cofactors . However, it has been shown thatsubstitution of a single amino acid alters GFOR from Z . mobilisto a glucose dehydrogenase with dual cofactor specificity forNADP⁺ and NAD⁺ [42] . Thus, one might speculate that H . marismortuixylose dehydrogenase represents a natural mutant of an aldose-ketoseoxidoreductase.

 
The phylogenetic relationship of the xylose dehydrogenase ofH . marismortui with oxidoreductase and dehydrogenase sequences, showing significant similarity according to BLASTP searches[see above], is given in the phylogram shown in Fig . 7 . They include the characterized GFOR from Z . mobilis and putative bacterial GFORs [cluster IA], putative archaeal oxidoreductases and dehydrogenases [IB], eucaryal xylose dehydrogenases-dimericDDs [II], and archaeal glucose dehydrogenases [III], each forminga separate cluster . In accordance with the highest degree of similarity, the H . marismortui sequence clusters within the bacterial oxidoreductases for which only the GFOR from Z . mobilis has been functionally characterized . Cluster IB contains only putative archaeal oxidoreductases and dehydrogenases, including dehydrogenases from Sulfolobus and Pyrococcus spp . Determination of whether the putative bacterial or archaeal sequences of clusters IA and IB represent functional oxidoreductases or [xylose] dehydrogenases must await their biochemical characterization following expression of the genes that encode them . Eukaryotic xylose dehydrogenases-dimeric DDs, [cluster II], which probably represent a novel family of dehydrogenases [1, 2], and archaeal glucose dehydrogenases [clusterIII], which belong to the medium-chain dehydrogenase-reductasesuperfamily [26], form distinct phylogenetic clusters separatefrom the xylose dehydrogenase from H . marismortui, which isin accordance with differences in their molecular and catalyticproperties.

Is xylose dehydrogenase the first enzyme of archaeal xylose catabolism? The xylose dehydrogenase of H . marismortui showed specific inductionduring growth on xylose; together with the findings that xyloseisomerase and xylulose kinase, as well as the genes that encodethem, were absent from H . marismortui, we suggest that thisnovel type of xylose dehydrogenase represents the first stepin xylose degradation by this archaeon . Since xylose isomeraseand xylulose kinase and the genes that encode them, xylA andxylB, have not been reported for any archaeal species, one mightspeculate that the initial reaction of xylose metabolism in archaea in general might involve a xylose dehydrogenase rather than xylose isomerase and xylulose kinase, the typical reactionsin bacterial xylose catabolism . Further steps in the xylosedegradation pathway in H . marismortui following the fate ofxylonate remain to be elucidated . Experiments using a proteomicapproach to identify xylose-inducible proteins as analyzed bytwo-dimensional gel electrophoresis are in progress.

 
We thank R . Schmid [Mikrobiologie, Universität Osnabrück,Osnabrück, Germany] for performing N-terminal amino acidsequencing . We thank Shiladitya DasSarma for getting accessto available contigs of the genome of H . marismortui [NationalScience Foundation grant reference MCB-0135595; University ofMaryland Biotechnology Institute website [http://zdna2.umbi.umd.edu]]. The expert technical assistance of A . Brandenburger is gratefully acknowledged.

This work was supported by the EU grant Extremophiles as Cell Factories and by the Fonds der Chemischen Industrie.

 
* Corresponding author . Mailing address: Institut für Allgemeine Mikrobiologie, Christian-Albrechts-Universität Kiel, Am Botanischen Garten 1-9, D-24118 Kiel, Germany . Phone: 49-431-880-4328 or 4330 . Fax: 49-431-880-2194 . E-mail: peter.schoenheit@ifam.uni-kiel.de .

Dedicated to Rolf Thauer on the occasion of his 65th birthday.

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