novel xylose dehydrogenase in the halophilic archaeon ... · larity to glucose-fructose...

JOURNAL OF BACTERIOLOGY, Sept. 2004, p. 6198–6207 Vol. 186, No. 180021-9193/04/$08.00�0 DOI: 10.1128/JB.186.18.6198–6207.2004Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Novel Xylose Dehydrogenase in the Halophilic ArchaeonHaloarcula marismortui†

Ulrike Johnsen and Peter Schonheit*Institut fur Allgemeine Mikrobiologie, Christian-Albrechts-Universitat 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 dehydro-genase was induced. The enzyme was purified to homogeneity. It constitutes a homotetramer of about 175 kDaand catalyzed the oxidation of xylose with both NADP� and NAD� as cosubstrates with 10-fold higher affinityfor NADP�. In addition to D-xylose, D-ribose was oxidized at similar kinetic constants, whereas D-glucose wasused with about 70-fold lower catalytic efficiency (kcat/Km). With the N-terminal amino acid sequence of thesubunit, an open reading frame (ORF)—coding for a 39.9-kDA protein—was identified in the partiallysequenced genome of H. marismortui. The function of the ORF as the gene designated xdh and coding for xylosedehydrogenase was proven by its functional overexpression in Escherichia coli. The recombinant enzyme wasreactivated from inclusion bodies following solubilization in urea and refolding in the presence of salts,reduced and oxidized glutathione, and substrates. Xylose dehydrogenase showed the highest sequence simi-larity to glucose-fructose oxidoreductase from Zymomonas mobilis and other putative bacterial and archaealoxidoreductases. Activities of xylose isomerase and xylulose kinase, the initial reactions of xylose catabolism ofmost bacteria, could not be detected in xylose-grown cells of H. marismortui, and the genes that encode them,xylA and xylB, were not found in the genome of H. marismortui. Thus, we propose that this first characterizedarchaeal xylose dehydrogenase catalyzes the initial step in xylose degradation by H. marismortui.

The utilization of sugars, in particular of hexoses and hexosepolymers and—to a lesser extent—of pentoses, has been re-ported for various species in the domain Archaea. So far, onlythe catabolic pathways of hexoses and glucose polymers (e.g.,maltose and starch) have been studied in detail in particularin hyperthermophilic, thermoacidophilic, and extremely halo-philic archaea. Comparative analyses of glucose degradationpathways in these organisms revealed that the classical Emb-den-Meyerhof- (EM) or Entner-Doudoroff- (ED) pathwayfound in bacteria is not operative in archaea; they use insteadmodified versions of these pathways as follows (for reviews, seereferences 31 and 41). In hyperthermophilic eury- and cren-archaeota, glucose degradation proceeds predominantly viamodified EM pathways, which differ from the classical EMpathway by the presence of several unusual glucokinases (ADPor ATP dependent) and 6-phosphofructokinases (ADP, ATP,or PPi dependent), novel enzymes of glucose-6-phosphateisomerization and of glyceraldehyde-3-phosphate oxidation,and pyruvate kinases with reduced regulatory potential (15,18, 41).

In thermoacidophilic archaea, Sulfolobus and Thermoplasmaspp., glucose is degraded via a nonphosphorylated version ofthe ED pathway (22, 31, 41) by which glucose is oxidized toglycerate via the nonphosphorylated intermediates gluconateand 2-keto-3-deoxygluconate (KDG) involving glucose dehy-drogenase, gluconate dehydratase, and KDG aldolase. Glycer-ate is then phosphorylated via a specific kinase to 2-phospho-glycerate, which is further converted to pyruvate via enolase

and pyruvate kinase. In halophilic archaea, e.g., Halococcus,Haloarcula, and Haloferax spp., a modified, semiphosphory-lated ED pathway is operative in which—as in thermoacido-philes—glucose is converted to KDG. However, KDG is thenphosphorylated to 2-keto-3-deoxy-6-phosphogluconate by KDGkinase. Further degradation of 2-keto-3-deoxy-6-phosphoglu-conate proceeds via reactions of the conventional phosphory-lated ED pathway found in bacteria (19, 45).

In contrast to hexose metabolism, the catabolic pathwaysof pentoses have not been studied in detail in the domainArchaea. The utilization of pentoses, e.g., xylose, ribose, andarabinose, has been reported for several halophiles, e.g., Halo-coccus, Haloarcula, and Halobacterium spp., and for Sulfolobusspecies (30, 40), rather than for the majority of hyperthermo-philes. No studies of growth on pentoses or analyses of theenzymes involved in pentose degradation by these organismshave been reported.

In the domain Bacteria, the pathways for the degradation ofpentoses, in particular, D-xylose, have been studied in detail inmany species, including Escherichia coli, Salmonella entericaserovar Typhimurium, Lactobacillus pentosus, Lactococcus lac-tis, Bacillus spp., Staphylococcus xylosus, Bacteroides xylanolyti-cus, and Tetragenococcus halophilus. Degradation of xylose bythese organisms, e.g., by E. coli, starts with its uptake via spe-cific high- or low-affinity transport systems. Via xylose isomer-ase, xylose is then isomerized to xylulose, which is phosphor-ylated to xylulose-5-phosphate by the activity of xylulosekinase. The genes encoding xylose transporters, xylose isomer-ase (xylA gene), and xylulose kinase (xylB gene), which are ar-ranged in an operon, are induced by xylose mediated by thetranscriptional regulator XylR. Further degradation of xylu-lose-5-phosphate, proceeds—depending on the organism—ei-ther via the pentose phosphate cycle, the phosphoketolase

* Corresponding author. Mailing address: Institut fur AllgemeineMikrobiologie, Christian-Albrechts-Universitat Kiel, Am BotanischenGarten 1-9, D-24118 Kiel, Germany. Phone: 49-431-880-4328 or 4330.Fax: 49-431-880-2194. E-mail: [email protected].

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

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pathway, or—as in Bacteroides spp. (4)—via a combination ofboth pentose phosphate and the EM pathway. Thus, the mostcommon initial reactions of bacterial xylose catabolism involvexylose isomerase and xylulose kinase (4, 13, 23, 29, 36).

Xylose isomerase and the gene that encodes it, xylA, havebeen characterized in many bacteria, including the hyperther-mophile Thermotoga maritima, as well as in eucarya. The en-zymes from both domains are of significant industrial interestsince they also catalyze, as a side activity, the isomerization ofglucose to fructose, a reaction that constitutes the last step inthe large-scale industrial process of the production of sweet-eners from starch (3).

So far, activities of xylose isomerase and xylulose kinase havenot been reported in any species of the archaeal domain. Fur-ther, homologs to the bacterial xylA and xylB genes could notbe found in archaeal genomes. Thus, it might be speculatedthat the initial steps in xylose degradation by archaea are dif-ferent 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. Evi-dence is presented that the first step in xylose degradation bythis organism is oxidation of xylose to xylonate via a xylose-inducible NADP�-reducing D-xylose dehydrogenase. This firstarchaeal xylose dehydrogenase was purified, and the gene thatencodes it, xdh, was identified in available sequenced contigs ofH. marismortui. This enzyme represents a novel type of xylosedehydrogenase, showing high similarity to glucose-fructose ox-idoreductase (GFOR) from Zymomonas mobilis.

MATERIALS AND METHODS

Growth of H. marismortui and preparation of cell extracts. H. marismortui(DSM 3752) (24) was obtained from the Deutsche Sammlung von Mikroorgan-ismen und Zellkulturen (Braunschweig, Germany). The organism was grownaerobically at 37°C in 500-ml Erlenmeyer flasks filled with 50 ml of mediumcontaining 25 mM xylose, 0.05% yeast extract, 250 g of NaCl per liter, 20 g ofMgSO4 � 7H2O per liter, 19.5 g of morpholineethanesulfonic acid (MES) perliter, 2 g of KCl per liter, 1 g of Na-glutamate per liter, and 3 g of Na-citrate perliter; 10 ml of the vitamin solution described by Staley (35); and 10 ml of a traceelement 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 ofZnSO4, and 0.01 g of CuSO4 � 5H2O. The pH was adjusted to 7.35 with 10 NNaOH. Growth was followed by measuring optical density at 578 nm (�OD578).During growth, samples were removed and centrifuged and the supernatantswere analyzed for xylose and xylonate, as indicated. Extracts were prepared fromlate-log-phase cells by sonication as described previously (19), and enzyme ac-tivities were determined as described in the section on enzyme assays. Protein wasdetermined by the biuret method with bovine serum albumin as the standard (5).

Induction of xylose dehydrogenase in H. marismortui was followed in 2,000-mlErlenmeyer flasks filled with 400 ml of medium containing xylose (25 mM), yeastextract (2.5 g/liter), and Casamino Acids (5 g/liter); cells previously grown onglucose (see reference 19) were used as an inoculum (10%). At the timesindicated, 60- to 80-ml samples were removed and centrifuged (2,600 � g, 10min, 4°C) and the cell pellets were suspended in 1 ml of 0.1 M Tris-HCl, pH 7.5,containing 250 g of NaCl/liter. Cell extracts were prepared by sonication (19),followed by centrifugation at 12,000 � g for 10 min. The supernatants wereanalyzed for xylose dehydrogenase activities. The protein concentration of cellextracts was determined by the biuret method.

Purification of xylose dehydrogenase from H. marismortui. Xylose dehydroge-nase was purified from H. marismortui after growth of the organism in a medium(see above) containing 25 mM xylose and 0.1% yeast extract in a 10-liter Biostatfermentor. Extract was prepared from 4 g of cells, which were suspended in 100mM Tris-HCl, pH 8.8, containing 2 M (NH4)2SO4 and 20 mM MgCl2 (buffer A).Cells were disrupted by passage through a French pressure cell at 1.3 � 108 Pa.Cell debris and unbroken cells were removed by centrifugation for 90 min at100,000 � g at 4°C. The 100,000 � g supernatant was applied to a Sepharose CL4B column (1.6 by 60 cm) that had been equilibrated with buffer A. Protein was

eluted with a decreasing (NH4)2SO4 gradient from 2 to 0 M in buffer A. Frac-tions containing the highest xylose dehydrogenase activity [1.6 to 1.4 M(NH4)2SO4] were pooled, adjusted to 2 M (NH4)2SO4, and applied to a PhenylSepharose column (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 was eluted witha linear gradient of buffer B to 50 mM Tris-HCl, pH 8.5, containing 20 mMMgCl2 and 10% glycerol. Fractions containing the highest xylose dehydrogenaseactivity [1.04 to 0.95 M (NH4)2SO4] were pooled and concentrated to 600 �l byultrafiltration (cutoff, 20 kDa). The concentrated protein solution was applied toa Superdex 200 HiLoad gel filtration column (1.6 by 60 cm) that had beenequilibrated with 50 mM Tris-HCl, pH 8.5, containing 20 mM MgCl2, 10%glycerol, and 100 mM NaCl. In this buffer, the enzyme was stable in the absenceof a high salt (KCl or NaCl) concentration. Eluted fractions containing xylosedehydrogenase activity indicated essentially pure protein and were stored at�20°C. The purity of the preparations was checked by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) in 12% gels in accordance withstandard procedures (21). During the purification procedure, protein concentra-tions were determined by the Bradford method with bovine serum albumin as thestandard (7).

Cloning and expression of xylose dehydrogenase from H. marismortui inE. coli. On the basis of the N-terminal amino acid sequence, an open readingframe (ORF) was identified by a BLASTP search in contig 97 of the partiallysequenced genome of H. marismortui (P. Zhang, W. V. Ng, and S. DasSarma,personal communication, 2003). The ORF was characterized as the xdh gene,encoding xylose dehydrogenase, by its functional overexpression in E. coli. Thegene was amplified from genomic DNA of H. marismortui by PCR and clonedinto pET17b (Novagen) via two restriction sites (NdeI and BamHI) created withthe primers 5�-GACGACAGTCATATGAACGTTG-3� and 5�-CAAAAAATCTGGATCCGGTTTC-3� (restriction sites are underlined). The vector pET17b-xdh was transformed into E. coli BL21 codon plus(DE3)-RIL (Stratagene). Forexpression, cells were grown in Luria-Bertani medium at 37°C. Expression wasinitiated by the addition of isopropyl-�-D-thiogalactopyranoside (IPTG; finalconcentration, 0.4 mM). After 18 h of further growth, cells were harvested bycentrifugation.

Solubilization, refolding, and purification of recombinant xylose dehydroge-nase. Recombinant xylose dehydrogenase, which was expressed in inclusionbodies, was solubilized and refolded as described by Connaris et al. (11), withmodifications. The E. coli cell pellets were suspended in 20 mM Tris-HCl, pH7.5, containing 2 mM EDTA, 3 M KCl, and 10% glycerol (buffer C). The cellsuspension was treated with 100 �g of lysozyme per ml and 0.1% (vol/vol) TritonX-100 and incubated at 30°C for 60 min, followed by incubation on ice for 15min. The suspension was then sonicated and centrifuged at 40,000 � g for 30 minat 4°C. The insoluble fraction was washed twice in buffer C, yielding inclusionbodies and insoluble cell fragments. The insoluble fraction was dissolved in 20mM Tris-HCl, pH 7.5, containing 2 mM EDTA, 8 M urea, and 50 mM dithio-erythritol. Solubilization was carried out at 37°C for 1 h. Refolding was initiatedby slowly diluting the suspension in 20 mM Tris-HCl, pH 7.5, containing 3 MKCl, 2 mM EDTA, 10% glycerol, 2 mM xylose, 0.1 mM NADP�, 3 mM reducedglutathione, and 0.3 mM oxidized glutathione to a final concentration of about 30�g of protein per ml. After incubation for 6 days at 4°C, the renatured proteinwas concentrated 250-fold by ultrafiltration (cutoff, 30 kDa). The concentratedprotein solution was applied to a Superdex 200 HiLoad 16/60 gel filtrationcolumn that had been equilibrated with 50 mM Tris-HCl, pH 8.5, containing 20mM MgCl2, 10% glycerol, and 100 mM NaCl. Eluted fractions containing xylosedehydrogenase activity were pooled and applied to a Phenyl Resource column (1ml), equilibrated with buffer B. Protein was eluted with a linear (NH4)2SO4

gradient to 0 M in 50 mM Tris-HCl, pH 8.5, containing 20 mM MgCl2 and 10%glycerol. Essentially pure enzyme was eluted at about 1 M (NH4)2SO4. Thepurity of the preparations was checked by SDS-PAGE, and protein concentra-tions 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 �mol of substrate consumed orproduct formed per min.

Xylose dehydrogenase activity (xylose � NADP� 3 xylonate � NADPH �H�) was assayed by measuring the rate of reduction of NADP� at 365 nm. Thestandard assay mixture contained 100 mM Tris-HCl (pH 8.3), 1.5 M KCl, 1 mMNADP�, 10 mM xylose, and protein.

Glucose dehydrogenase activity was tested as glucose-dependent reduction ofNADP�. The assay mixture contained 100 mM Tris-HCl (pH 8.3), 1.5 M KCl, 10mM glucose, 1 mM NADP�, and protein.

Xylose isomerase activity was tested as xylose-dependent formation of xylu-lose. The assay mixture contained 100 mM Tris-HCl (pH 8.5), 1 M KCl, 1 mMCoCl2, 5 mM MnSO4, 5 mM MgCl2, 10 to 100 mM xylose or glucose, and protein.

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During incubation (0 to 20 min), aliquots were taken and the reaction wasstopped by addition of trichloroacetic acid to a final concentration of 10%. Aftercentrifugation, xylulose was quantified by the cysteine-carbazole method (17).Crude extract from xylose-grown E. coli cells served as a positive control (12).

Xylulose-5-phosphate kinase activity was tested as the ATP-dependent de-crease in xylulose. The assay mixture contained 100 mM Tris-HCl (pH 8.5), 1 MKCl, 10 mM MgCl2, 4 mM cysteine-HCl, 10 mM ATP, 6 mM xylulose, andprotein. During incubation (0 to 30 min), aliquots were taken and the reactionwas stopped by addition of 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. Thetemperature dependence of xylose dehydrogenase was measured between 20 and60°C in 50 mM Tris-HCl, pH 8.3, containing 1.5 M KCl, 1 mM NADP�, 10 mMxylose, and protein. The pH dependence of the enzyme was measured betweenpHs 4.4 and 10.3 at 37°C with either piperazine (pHs 4.9 to 6.0), bis-Tris (pHs 6.0to 7.5), 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 ofsalts (0 to 200 mM MgCl2, 0 to 3.5 M KCl, and 0 to 3.5 M NaCl) on xylosedehydrogenase activity were tested at 37°C in 20 mM Tris-HCl, pH 8.3, contain-ing 1 mM NADP�, 10 mM xylose, and protein.

Substrate specificity. The substrate specificity of xylose dehydrogenase wastested at 37°C in 20 mM Tris-HCl, pH 8.3, containing 1.5 M KCl with D isomersof the sugars in the presence of NADP� at 10 mM each xylose and ribose; 1 mMNADP�; 100 mM glucose, 1 mM NADP�; 100 mM each galactose, fructose, andarabinose; and 2 mM NADP�. For the determination of apparent Km and Vmax

values for sugars and the cosubstrates NADP� and NAD�, the following con-centrations were used: xylose or ribose, 0 to 10 mM with 1 mM NADP�;NADP�, 0 to 1 mM with 10 mM xylose or ribose; NAD�, 0 to 3 mM with 10 mMxylose or ribose; glucose, 0 to 100 mM with 1 mM NADP�; NADP�, 0 to 1 mMwith 100 mM glucose.

Analytical assays. Gel filtration chromatography was carried out with a flowrate of 1 ml/min on a Superdex 200 HiLoad column (1.6 by 60 cm). The columnwas equilibrated with 50 mM Tris-HCl, pH 8.5, containing 20 mM MgCl2, 10%glycerol, and 100 mM NaCl. HWM and LWM gel filtration calibration kits(Amersham Biosciences, Amersham, England) were used as the standards.

The concentration of xylose was determined by using the orcinol assay (8). Theextinction coefficient (ε) at 546 nm was 4,800 M�1 cm�1. The concentration ofxylonate was determined by high-performance liquid chromatography with anAminex HPX87H column (Bio-Rad, Richmond, Calif.) operating at 37°C. Sam-ples were diluted 1:5 in 5 mM H2SO4, boiled for 30 min, centrifuged, passedthrough a 0.2-�m-pore-size filter, and loaded onto the column. Xylonate waseluted with 5 mM H2SO4 at a flow rate of 0.6 ml/min and then monitored witha differential refractometer at 210 nm.

RESULTS

Growth of H. marismortui on xylose and induction of xylosedehydrogenase. H. marismortui was grown on a medium con-taining 25 mM D-xylose and 0.05% yeast extract, with xylose-grown cells (10%) as the inoculum. The cells grew exponen-tially with a doubling time of about 20 h up to a �OD578 ofabout 1. During growth, xylose was consumed and smallamounts of xylonate were formed (Fig. 1). In the absence ofxylose, the cells grew with a doubling time of about 30 h up toa �OD578 of about 0.5 because of the yeast extract present inthe medium (data not shown).

To identify the first enzymes of xylose degradation, extractsof xylose-grown H. marismortui cells were analyzed for xyloseisomerase and xylulose kinase, the initial enzymes of xylosedegradation by bacteria. Neither of these activities could bedetected. As a control, both enzyme activities (xylose isomer-ase, 40 mU/mg; xylulose kinase, 420 mU/mg) were found inxylose-grown E. coli cells under conditions identical to thoseused for H. marismortui.

Since during growth on xylose small amounts of xylonatewere formed, we looked for enzymes catalyzing the dehydro-genation of xylose. Indeed, extracts of xylose-grown cells cat-

alyzed the NADP�-dependent conversion of xylose to xylonateat a specific activity of 0.15 U/mg with an apparent Km forxylose of 0.95 mM. Extracts of xylose-grown cells also catalyzedthe oxidation of glucose with NADP� to gluconate, however,at a 70-fold lower catalytic efficiency (apparent Vmax, 0.03 U/mg; Km, 15 mM), indicating that xylose-grown H. marismortuicells contain a specific xylose dehydrogenase different fromglucose dehydrogenase. Glucose-grown cells of H. marismortuialso contained xylose dehydrogenase activity, however, withabout 10-fold lower catalytic efficiency (apparent Vmax, 0.03 U/mg; Km, 2 mM) compared to that of xylose-grown cells, sug-gesting that xylose dehydrogenase was induced during growthon xylose.

Induction of xylose dehydrogenase was demonstrated duringgrowth of H. marismortui on xylose with cells pregrown onglucose as an inoculum. Because of the higher concentrationsof yeast extracts and Casamino Acids present in the medium,the cells grew with a shorter doubling time (about 12 h) tosignificantly higher cell densities and the amount of xyloseconsumed increased (Fig. 2). During growth, the NADP�-dependent xylose dehydrogenase activity increased up to five-fold parallel to xylose consumption, indicating that the enzymeis induced by xylose and probably represents the first reactionof 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 chromatographicsteps. The most efficient purification step was hydrophobicinteraction chromatography on Phenyl Sepharose, resulting in130-fold enrichment. With the entire procedure, the enzymewas purified about 210-fold, to a specific activity of 100 U/mgwith a yield of 10% (Table 1). The purified protein was elec-trophoretically homogeneous as judged by denaturing SDS-

FIG. 1. Growth of H. marismortui on 25 mM xylose and 0.05%yeast extract. Cultures were incubated at 37°C in 500-ml Erlenmeyerflasks filled with 50 ml of medium and shaken at 200 rpm. Growth (■),xylose consumption (F), and xylonate formation (Œ) were followedover time.

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PAGE (Fig. 3). Thus, xylose dehydrogenase represents about0.5% of the cellular protein of H. marismortui.

Molecular and catalytic properties. The apparent molecularmass of native xylose dehydrogenase determined by gel filtra-tion on Superdex 200 was 175 � 15 kDa. SDS-PAGE revealedonly one subunit with an apparent molecular mass of 57 � 3kDa (Fig. 3). This value is significantly overestimated, as hasbeen observed for various halophilic enzymes (see Discussion).Recombinant xylose dehydrogenase (see below), showed an ap-parent molecular mass on SDS-PAGE of 57 � 3 kDa, althoughthe calculated molecular mass is 39.9 kDa. We propose thatnative xylose dehydrogenase is a homotetrameric (4) enzyme.

The purified enzyme catalyzed the oxidation of D-xylose withboth NADP� and NAD�. The rate dependence of the enzymeon xylose, NADP�, and NAD� followed Michaelis-Mentenkinetics with apparent Km values of 1.2, 0.15, and 0.9 mM,respectively. The corresponding Vmax values were about 100,

92, and 80 U/mg. The sixfold higher apparent Km for NAD�

compared to NADP� indicates that NADP� is the preferredelectron acceptor. In addition to D-xylose, various other pen-toses and hexoses (all D isomers) were tested as substrates forthe dehydrogenase with NADP� as the electron acceptor. Theapparent Km values, Vmax, kcat values, and catalytic efficiencies(kcat/Km) are given in Table 2. The highest catalytic efficiencywas obtained with xylose and NADP�. D-Ribose was also ac-cepted by the enzyme at high efficiency, whereas D-arabinosewas not oxidized at significant rates, indicating that the con-figuration change at C-2 significantly affects enzyme activity.

FIG. 2. Induction of xylose dehydrogenase during growth of H. marismortui on xylose. A culture was incubated at 37°C in a 2,000-ml Erlenmeyerflask filled with 400 ml of medium containing xylose (25 mM), yeast extract (2.5 g/liter), and Casamino Acids (5 g/liter) and shaken at 200 rpm.Glucose-grown cells were used as the inoculum (10%). Growth (■), xylose consumption (F), and the specific activity of xylose dehydrogenase (Œ)were followed over time. Protein concentration was determined by the biuret method.

FIG. 3. Purified xylose dehydrogenase from H. marismortui (A) andrecombinant xylose dehydrogenase from transformed E. coli (B) as ana-lyzed by SDS-PAGE. (A) Lanes: 1, molecular mass standards; 2, nativeenzyme purified from H. marismortui. (B) Lanes: 1, molecular mass stan-dards; 2, recombinant enzyme purified from E. coli.

TABLE 1. Purification of xylose dehydrogenasefrom H. marismortui

Purification stepAmt ofproteinb

(mg)

Enzymeactivitya

(U)

Sp act(U/mg)

Yield(%)

Purifi-cation(fold)

Cell extract 260 125 0.48 100 1Sepharose CL 4B 24 102 4.4 82 9Phenyl Sepharose 0.57 37 66 30 137Superdex 0.12 12 100 10 211

a Enzyme activity was measured at 37°C as NADP� and xylose-dependentxylonate formation. During the purification procedure, the assay mixture con-tained 100 mM Tris-HCl (pH 8.8), 1.5 M KCl, 1 mM NADP�, and 10 mM xylose.

b Protein concentration was determined by the Bradford method (7).



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D-Glucose was oxidized with a catalytic efficiency 70-fold lowerthan that with which xylose was oxidized, thus defining theenzyme as a specific xylose dehydrogenase (Fig. 4 A and B).Galactose oxidation was less efficient than glucose oxidation,and almost no activity was found with fructose (Table 2).

Effects of salt, pH, and temperature on xylose dehydroge-nase activity. The activity of xylose dehydrogenase from H.marismortui was strongly stimulated by high concentrations ofNaCl or KCl and by moderate concentrations of MgCl2. Max-imal activities were obtained at about 1.5 M both KCl and

NaCl and at 100 mM MgCl2 (Fig. 5). The rate dependence ofthe enzyme on pH and temperature was tested in the presenceof 1.5 M KCl. The pH optimum was about pH 8.3, and 50%activity was found at pHs 7 and 9. Xylose dehydrogenase ac-tivity increased exponentially with temperature between 25 and45°C; from the corresponding linear part of an Arrhenius plot,an activation energy of 64 kJ/mol was calculated. The highestcatalytic activity of xylose dehydrogenase was found at 50°C.

Identification, sequence analysis, and cloning of the geneencoding xylose dehydrogenase from H. marismortui and itsfunctional overexpression in E. coli. On the basis of the N-ter-minal amino acid sequence determined from the subunit ofxylose dehydrogenase, MNVDALTGGFDRRDWQEQTATDNPVRFAA, an ORF that exactly matches the 29 N-terminalamino acid residues was identified by BLASTP search in contig97 of the partially sequenced genome of H. marismortui (Zhanget al., personal communication). The ORF contains 1,083 bpcoding for a polypeptide of 360 amino acids with a calculatedmolecular mass of 39.9 kDa; the protein contains large amountsof negatively charged amino acids, 10% Asp and 11% Glu,which is typical for halophilic enzymes (10, 27), and had apredicted pI of 4.2. The G�C content of the ORF is 62 mol%.The coding sequence starts with ATG and stops with TGA. Aputative archaeal box A (TATA box) promoter signal (5�-TAATAT-3�) was identified between positions �23 and �28 up-stream from the ATG start codon (25, 37). Immediately up-stream of the initiation codon, a putative ribosome binding sitewith the sequence 5�-GTGGT-3� is present (32). Downstreamof the gene, a pyrimidine-rich sequence beginning at position1069 and a short inverted repeat located between positions

FIG. 4. Rate dependence of xylose dehydrogenase purified from H. marismortui on the concentrations of xylose (A) and glucose (B). Theinserts show double-reciprocal plots of the rates versus the corresponding substrate concentrations. The assay mixture contained 20 mM Tris-HCl(pH 8.3), 1.5 M KCl, 1 mM NADP�, various concentrations of xylose or glucose, and enzyme.

TABLE 2. Kinetic parameters of purified xylose dehydrogenasefrom H. marismortuia

Substrate Vmax(U/mg) Km (mM) kcat (s�1) kcat/Km

(s�1, 102)

D-Xyloseb 107 1.2 356 2.97D-Ribosec 108 2.3 360 1.57D-Glucosed 82 64 273 0.043D-Galactosee 39 NDh

D-Fructosef 2 NDD-Arabinoseg 1 ND

a Enzyme activity was measured at 37°C with 0.1 M Tris-HCl (pH 8.3)–1.5 MKCl.

b Concentrations under Vmax conditions: D-xylose, 10 mM; NADP�, 1 mM.c Concentrations under Vmax conditions: D-ribose, 10 mM; NADP�, 1 mM.d Concentrations under Vmax conditions: D-glucose, 100 mM; NADP�, 1 mM.e Concentrations under Vmax conditions: D-galactose, 100 mM; NADP�,

2 mM.f Concentrations under Vmax conditions: D-fructose, 100 mM; NADP�,

2 mM.g Concentrations under Vmax conditions: D-arabinose, 100 mM; NADP�,

2 mM.h ND, not determined.



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1112 and 1121 were identified, indicating a transcription ter-mination site (38).

The ORF was characterized as the xdh gene encoding xylosedehydrogenase by its functional overexpression in E. coli. Thexdh gene was amplified by PCR and cloned into the vectorpET17b. The recombinant plasmid was used to transformE. coli BL21 codon plus(DE3)-RIL. After induction with IPTG,a polypeptide of about 59 kDa was overexpressed, which wasrecovered almost completely in inclusion bodies. The proteinpurified from inclusion bodies was a catalytically active, ex-tremely halophilic xylose dehydrogenase.

Solubilization, refolding, and purification of recombinantxylose dehydrogenase. Recombinant xylose dehydrogenase waspurified from inclusion bodies by dissolving with 8 M urea inthe presence of dithioerythritol, followed by refolding with abuffer containing a high salt concentration (3 M KCl), sub-strates, and glutathione (see Materials and Methods). Maximalxylose dehydrogenase activity was obtained after 6 days of in-cubation. The refolded activated xylose dehydrogenase was pu-rified by chromatography on Superdex and Phenyl Resource.The recombinant xylose dehydrogenase showed molecular andkinetic properties almost identical to those of the enzyme pu-rified from H. marismortui (Table 2). The molecular masses ofthe native enzyme and subunits were 180 � 10 and 59 � 3 kDa(Fig. 3), respectively, and the apparent Km values were very sim-ilar for xylose, ribose, glucose, and NADP� (data not shown);however, the apparent Vmax of the recombinant enzyme wassignificantly (about 40%) lower.

DISCUSSION

In the present communication, we describe the purificationand characterization of the first archaeal D-xylose dehydroge-nase and the gene that encodes it from the halophilic archaeonH. marismortui. The enzyme was induced during growth onxylose, suggesting that xylose dehydrogenase represents theinitial enzyme of the xylose degradation pathway in this ar-

chaeon. The enzyme constitutes a novel type of xylose dehy-drogenase related to GFOR from Z. mobilis.

Molecular and kinetic properties. Xylose dehydrogenasewas characterized as a homotetrameric enzyme of about 175kDa; the calculated subunit molecular mass is 39.9 kDa. Theapparent molecular mass of subunits on SDS-PAGE of about57 kDa obtained for xylose dehydrogenase was overestimatedas reported for several halophilic proteins, probably because ofthe presence of large amounts of negatively charged aminoacids. The same degree of overestimation as described for xy-lose dehydrogenase was observed with glucose dehydrogenasefrom Haloferax mediterranei. The enzyme has a calculated mo-lecular mass of 39.3 kDa (27) and showed an apparent molec-ular mass on SDS-PAGE of 53 � 3 kDa (6).

Xylose dehydrogenase showed dual cofactor specificity forpyridine nucleotides with a high preference for NADP� overNAD�, indicating that NADP� is the physiological electronacceptor. The enzyme catalyzed the oxidation of xylose, ribose,and glucose; however, the catalytic activity for the latter wasabout 70-fold lower. The archaeal xylose dehydrogenase can bediscriminated from archaeal glucose dehydrogenases charac-terized from various organisms including Haloferax, Sulfolobus,Thermoplasma, and Thermoproteus spp., which all show xylosedehydrogenase activity. These archaeal glucose dehydrogenasesare tetrameric or dimeric enzymes composed of 40-kDa sub-units, show dual cofactor specificity for NADP� and NAD�

with a high preference for NADP�, and utilize various aldoses(hexose and pentoses) including xylose in addition to glucose.However, in contrast to xylose dehydrogenase from H. maris-mortui, all archaeal glucose dehydrogenases showed signif-icantly higher catalytic efficiencies for NADP�-dependentoxidation of glucose compared to that of xylose (6, 22, 33,34).

Few reports of purified xylose dehydrogenases from Eucaryaand Bacteria are available. An NADP�-specific xylose dehy-drogenase from pig liver was characterized (46). The enzyme is

FIG. 5. Effects of NaCl and KCl (A) and MgCl2 (B) on xylose dehydrogenase purified from H. marismortui. The assay mixture contained 20mM Tris-HCl (pH 8.3), 1 mM NADP�, 10 mM xylose, enzyme, and the concentrations of NaCl, KCl, and MgCl2 indicated.



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a homodimer composed of 32-kDA subunits showing the high-est catalytic activity with xylose but also accepts ribose andglucose at about 7- or 30-fold lower catalytic efficiency. Theenzyme was specific for NADP� and did not reduce NAD�.Recently, Aoki et al. (1) demonstrated that the NADP�-de-pendent D-xylose dehydrogenase of pig liver is identical to di-meric dihydrodiol dehydrogenase (DD). Copurification of DDactivity and xylose dehydrogenase activity from pig liver, mo-lecular mass and kinetic analyses, and inhibitor studies showedthat the two enzymes are identical (1). Dimeric DDs catalyzethe NADP�-dependent oxidation of various aromatic hydro-carbons, e.g., naphthalene dihydrodiol, to the correspondingcatechols and also the oxidation of various sugars, with xyloseas the most effective sugar substrate. The oxidation rate ofnaphthalene dihydrodiol was about twofold higher than that ofxylose (1), suggesting that dihydrodiols are the preferred sub-strates of the dimeric DD-xylose dehydrogenase. The genescoding for various mammalian dimeric DD-xylose dehydroge-nases, including pig liver, rabbit lens, human intestine, andmonkey kidney, were sequenced (2), and thus, sequences ofeucaryal xylose dehydrogenases are known.

In bacteria, oxidation of xylose to xylonate has been re-ported for several species (9); an NAD�-reducing xylose de-hydrogenase activity was reported for two Caulobacter species(28); however—to our knowledge—the only xylose dehydroge-nase from bacteria characterized to some detail is the enzymefrom Arthrobacter sp. The enzyme was induced by xylose andshowed a high specificity for xylose (apparent Km, 17.4 mM)and for NAD� as an electron acceptor. Other pentoses andhexoses, as well as NADP� as a cofactor, were not accepted assubstrates (44). The amino acid sequence of the enzyme hasnot been reported.

Thus, both characterized eucaryal and bacterial D-xylose de-hydrogenases showed significant differences in molecular andkinetic properties compared to the archaeal xylose dehydroge-nase from H. marismortui.

Sequence comparison and phylogenetic analysis of archaealxylose dehydrogenase. On the basis of the N-terminal aminoacid sequence of the subunit, the xdh gene encoding the xylosedehydrogenase from H. marismortui was identified in contig 97of the partially sequenced genome of the organism by func-tional overexpression in E. coli. BLASTP searches of nonre-dundant databases with the deduced amino acid sequence ofthe xdh gene from H. marismortui revealed various hits. Almostall of them were putative oxidoreductases or dehydrogenases.The highest degrees of similarity were found with bacterialGFOR from Z. mobilis (41%) and hypothetical GFORs fromthe bacteria Deinococcus radiodurans, Caulobacter crescentus,Streptococcus pneumoniae, and Bacillus halodurans (35 to40%). Similarities of 31 to 34% with the putative archaealoxidoreductases or dehydrogenases from Pyrococcus sp., Sul-folobus solfataricus, and Thermoplasma sp. were found, the bestscores being obtained for the putative dehydrogenases fromPyrococcus furiosus (PF1919) and S. solfataricus (SSO3015).Lower degrees of similarity were found with characterizedeucaryal xylose dehydrogenases-dimeric DDs (28 to 29%) andwith archaeal glucose dehydrogenases (17 to 21%) from halo-philes (Haloferax and Halobacterium spp.) and thermoacido-philes (Sulfolobus and Thermoplasma spp.). For accessionnumbers, see the legend to Fig. 7.

The most similar enzyme of xylose dehydrogenase from H.marismortui, i.e., GFOR from Z. mobilis, is a homotetramericenzyme composed of 40-kDa subunits and containing tightlybound cofactor NADP�. The enzyme catalyzes the coupledintermolecular oxidation-reduction of glucose and fructose toform gluconolactone and sorbitol. The periplasmic enzyme issynthesized as a precursor with an N-terminal signal peptide of52 amino acid residues (14). A multiple-sequence alignment ofxylose dehydrogenase of H. marismortui, mature GFOR fromZ. mobilis, and other putative bacterial oxidoreductases isgiven in Fig. 6. The alignment includes a prediction of second-ary structure, which is in accordance with the crystal structureof GFOR from Zymomonas oxidoreductase (20). Sequencecomparisons indicate a variety of conserved regions including atypical �� dinucleotide binding pocket (Rossman fold, aminoacids 26 to 57) (16, 43) and—with few deviations—the recentlypostulated consensus sequence for a novel class of dehydroge-nases, including the highly conserved EKP motif (42) (aminoacids 113 to 122). Although xylose dehydrogenase shows a highdegree of similarity to GFOR from Z. mobilis, the two enzymescatalyze different reactions; xylose dehydrogenase is a dehy-drogenase with dual cofactor specificity for NADP� andNAD�, whereas GFOR catalyzes the coupled oxidation-reduc-tion of glucose and fructose with tightly bound NADP�, i.e., inthe absence of added cofactors. However, it has been shownthat substitution of a single amino acid alters GFOR from Z.mobilis to a glucose dehydrogenase with dual cofactor speci-ficity for NADP� and NAD� (42). Thus, one might speculatethat H. marismortui xylose dehydrogenase represents a naturalmutant of an aldose-ketose oxidoreductase.

The phylogenetic relationship of the xylose dehydrogenaseof H. marismortui with oxidoreductase and dehydrogenase se-quences, showing significant similarity according to BLASTPsearches (see above), is given in the phylogram shown in Fig. 7.They include the characterized GFOR from Z. mobilis andputative bacterial GFORs (cluster IA), putative archaeal oxi-doreductases and dehydrogenases (IB), eucaryal xylose dehy-drogenases-dimeric DDs (II), and archaeal glucose dehydro-genases (III), each forming a separate cluster. In accordancewith the highest degree of similarity, the H. marismortui se-quence clusters within the bacterial oxidoreductases for whichonly the GFOR from Z. mobilis has been functionally charac-terized. Cluster IB contains only putative archaeal oxidoreduc-tases and dehydrogenases, including dehydrogenases from Sul-folobus and Pyrococcus spp. Determination of whether theputative bacterial or archaeal sequences of clusters IA and IBrepresent functional oxidoreductases or (xylose) dehydroge-nases must await their biochemical characterization followingexpression of the genes that encode them. Eukaryotic xylosedehydrogenases-dimeric DDs, (cluster II), which probably rep-resent a novel family of dehydrogenases (1, 2), and archaealglucose dehydrogenases (cluster III), which belong to the me-dium-chain dehydrogenase-reductase superfamily (26), formdistinct phylogenetic clusters separate from the xylose dehy-drogenase from H. marismortui, which is in accordance withdifferences in their molecular and catalytic properties.

Is xylose dehydrogenase the first enzyme of archaeal xylosecatabolism? The xylose dehydrogenase of H. marismortuishowed specific induction during growth on xylose; togetherwith the findings that xylose isomerase and xylulose kinase, as



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FIG. 6. Multiple-sequence alignment of deduced amino acid sequences of the xylose dehydrogenase from H. marismortui, of GFOR from Z.mobilis, and of putative bacterial oxidoreductases. Characterized proteins are marked by asterisks. The alignment was generated by ClustalX withthe Gonnet matrix (39). The predicted secondary structure of the xylose dehydrogenase from H. marismortui is shown above the sequences.Symbols denote residues of the Rossman fingerprint motif (16, 43): ‚, basic or hydrophilic; E, small and hydrophobic; F, glycine; —, acid. NationalCenter for Biotechnology Information accession numbers: C. crescentus, AAK23207; D. radiodurans, B75475; Xanthomonas axonopodis,AAM35776; X. campestris, AAM40130; Z. mobilis GFOR, 1H6DK.

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well as the genes that encode them, were absent from H.marismortui, we suggest that this novel type of xylose dehydro-genase represents the first step in xylose degradation by thisarchaeon. Since xylose isomerase and xylulose kinase and thegenes that encode them, xylA and xylB, have not been reportedfor any archaeal species, one might speculate that the initialreaction of xylose metabolism in archaea in general mightinvolve a xylose dehydrogenase rather than xylose isomeraseand xylulose kinase, the typical reactions in bacterial xylosecatabolism. Further steps in the xylose degradation pathway inH. marismortui following the fate of xylonate remain to be

elucidated. Experiments using a proteomic approach to iden-tify xylose-inducible proteins as analyzed by two-dimensionalgel electrophoresis are in progress.

ACKNOWLEDGMENTS

We thank R. Schmid (Mikrobiologie, Universitat Osnabruck, Os-nabruck, Germany) for performing N-terminal amino acid sequencing.We thank Shiladitya DasSarma for getting access to available contigsof the genome of H. marismortui (National Science Foundation grantreference MCB-0135595; University of Maryland Biotechnology Insti-tute website [http://zdna2.umbi.umd.edu]). The expert technical assis-tance of A. Brandenburger is gratefully acknowledged.

FIG. 7. Phylogenetic relationships of the archaeal xylose dehydrogenase from H. marismortui, oxidoreductases and dehydrogenases frombacteria (IA) and archaea (IB), xylose dehydrogenases-dimeric DDs from eucarya (II), and glucose dehydrogenases from archaea (III). Thenumbers at the nodes are bootstrapping values according to neighbor joining (generated by using the neighbor-joining options of ClustalX).National Center for Biotechnology Information accession numbers: C. crescentus, AAK23207; D. radiodurans, B75475; dog DD, BAA83487;Halobacterium sp. NRC-1 glcdh, AAG18991; H. mediterranei glcdh, CAC4250; human DD, BAA83490; monkey DD, BAA83488; pig DD,BAA83486; Pyrococcus abyssi ORF PAB1139, B75025; P. furiosus ORF PF1919, AAL82043; rabbit DD, BAA83485; S. solfataricus ORF SSO3015,AAK43117; S. solfataricus dhg-1 (ORF SSO3003), AAK43106; S. solfataricus dhg-2 (ORF SSO3042), AAK43143; S. solfataricus dhg-3 (ORFSSO3204), AAK43301; Thermoplasma acidophilum ORF TA1182, CAC12307; T. acidophilum gdh, CAA42450; T. volcanium TVG1418453,BAB60511; X. campestris, AAM40130; X. axonopodis, AAM35776; Z. mobilis GFOR, 1H6DK.



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

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