Xanthine dehydrogenase from Pseudomonas putida 86: specificity, oxidation–reduction potentials of its redox-active centers, and first EPR characterization

Biochimica et Biophysica Acta 1544 (2001) 151^165 www.elsevier.com/locate/bba

Xanthine dehydrogenase from Pseudomonas putida 86: speci¢city, oxidation^reduction potentials of its redox-active centers, and ¢rst EPR characterization Katja Parschat a , Christoph Canne b , Ju«rgen Hu«ttermann b , Reinhard Kappl b , Susanne Fetzner a; * a

AG Mikrobiologie, Fachbereich 7, Carl von Ossietzky Universita«t Oldenburg, P.O. Box 2503, D-26111 Oldenburg, Germany b Fachrichtung Biophysik und Physikalische Grundlagen der Medizin, Universita«t des Saarlandes, Klinikum Geba«ude 76, D-66421 Homburg/Saar, Germany Received 8 December 1999; received in revised form 6 September 2000; accepted 6 September 2000

Abstract Xanthine dehydrogenase (XDH) from Pseudomonas putida 86, which was induced 65-fold by growth on hypoxanthine, was purified to homogeneity. It catalyzes the oxidation of hypoxanthine, xanthine, purine, and some aromatic aldehydes, using NAD‡ as the preferred electron acceptor. In the hypoxanthine:NAD‡ assay, the specific activity of purified XDH was 26.7 U (mg protein)31 . Its activity with ferricyanide and dioxygen was 58% and 4%, respectively, relative to the activity observed with NAD‡ . XDH from P. putida 86 consists of 91.0 kDa and 46.2 kDa subunits presumably forming an K4 L4 structure and contains the same set of redox-active centers as eukaryotic XDHs. After reduction of the enzyme with xanthine, electron paramagnetic resonance (EPR) signals of the neutral FAD semiquinone radical and the Mo(V) rapid signal were observed at 77 K. Resonances from FeSI and FeSII were detected at 15 K. Whereas the observable g factors for FeSII resemble those of other molybdenum hydroxylases, the FeSI center in contrast to most other known FeSI centers has nearly axial symmetry. The EPR features of the redox-active centers of P. putida XDH are very similar to those of eukaryotic XDHs/xanthine oxidases, suggesting that the environment of each center and their functionality are analogous in these enzymes. The midpoint potentials determined for the molybdenum, FeSI and FAD redox couples are close to each other and resemble those of the corresponding centers in eukaryotic XDHs. ß 2001 Elsevier Science B.V. All rights reserved. Keywords: Xanthine dehydrogenase; Molybdenum hydroxylase; N-Heteroaromatic compound; Aromatic aldehyde; Electron paramagnetic resonance spectroscopy; Redox potential

Abbreviations: bicine, N,N-bis(2-hydroxyethyl)glycine ; EPR, electron paramagnetic resonance; INT, 2-(p-iodophenyl)-3-(p-nitrophenyl)-5-phenyl-2H-tetrazolium chloride; MALDI-TOF-MS, matrix-assisted laser desorption/ionization-time of £ight-mass spectrometry; Mo-MCD, molybdenum-molybdopterin cytosine dinucleotide; Mo-MPT, molybdenum-molybdopterin ; NBT, 3,3P-(3,3P-dimethoxy-4,4Pbiphenylylene)-bis[2-(p-nitrophenyl)-5-phenyl-2H-tetrazolium chloride]; PAGE, polyacrylamide gel electrophoresis ; SDS, sodium dodecyl sulfate; TLC, thin layer chromatography; Tris, 2-amino-2-(hydroxymethyl)-1,3-propanediol ; XDH, xanthine dehydrogenase; XO, xanthine oxidase * Corresponding author. Fax: +49-441-798-3250; E-mail: [email protected] 0167-4838 / 01 / $ ^ see front matter ß 2001 Elsevier Science B.V. All rights reserved. PII: S 0 1 6 7 - 4 8 3 8 ( 0 0 ) 0 0 2 1 4 - 4

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1. Introduction Xanthine oxidoreductase catalyzes the oxidation of hypoxanthine and xanthine to urate. In the hydroxylation reaction, an oxygen atom derived from water is incorporated into the substrate, and electron equivalents are released. Electrons from hypoxanthine or xanthine are transferred either to NAD‡ or to molecular oxygen. Whereas xanthine dehydrogenases (XDHs) are characterized by high reactivity toward NAD‡ , but low reactivity toward O2 , xanthine oxidases (XOs) are highly active with O2 and show negligible reactivity toward NAD‡ . According to the structure of the molybdenum center in the active site, the XOs/XDHs are presumed to belong to the family of molybdenum hydroxylases (i.e. XO family) of the mononuclear molybdenum enzymes [1^6]. Whereas eukaryotic XOs/XDHs show a native molecular mass of about 300 kDa and consist of two identical subunits, each containing molybdenum-molybdopterin (Mo-MPT), two distinct [2Fe2S] centers (FeSI and FeSII), and FAD [1,7,8], XDHs from prokaryotic sources di¡er considerably with respect to their molecular masses, subunit compositions, and even with respect to their redox-active centers (Table 1). XDH from Pseudomonas putida Fu1, for example, is composed of two di¡erent subunits forming an K4 L4 structure, containing molybdenum and iron^sulfur centers, but possessing cytochrome b instead of a £avin [9]. XDHs from two purinolytic Clostridium species were reported to require selenium for activity [10,11]. P. putida 86 is able to utilize quinoline, hypoxanthine or xanthine as sole source of carbon, nitrogen, and energy [12,13]. When grown on quinoline, this strain synthesizes the molybdenum hydroxylase quinoline 2-oxidoreductase, which exhibits an K2 L2 Q2 structure and besides FeSI, FeSII and FAD contains a dinucleotide form of the pyranopterin molybdenum cofactor, namely Mo-MPT cytosine dinucleotide (Mo-MCD) [14^18]. Actually, bacterial molybdenum hydroxylases involved in the degradation of Nheteroaromatic compounds usually contain MoMCD [3], whereas XDHs from P. putida 86, Pseudomonas aeruginosa, Comamonas acidovorans and Rhodobacter capsulatus as well as eukaryotic XDHs/XOs possess the Mo-MPT cofactor ([19,20], Table 1).

However, the XDH from Veillonella atypica contains Mo-MCD and is a heterotrimer as many molybdenum hydroxylases [21], and XDH from Eubacterium barkeri was suggested to contain a dinucleotide-type pyranopterin molybdenum cofactor [22] (Table 1). A distinct feature of XO from cow's milk and some other XOs/XDHs is their broad substrate speci¢city toward N-heteroaromatic compounds and even some aldehydes [1,23^25]. Here we report on the substrate and electron acceptor speci¢city of the prokaryotic XDH from P. putida 86 and compare it with other XDHs. Since the paramagnetic molybdenum(V) (Mo(V)) has been discovered in milk XO by Bray et al. in 1959 [26], electron paramagnetic resonance (EPR) spectroscopy has been employed extensively to study the redox-active centers of eukaryotic XOs/XDHs [1,27^33]. EPR data on prokaryotic XDHs are comparatively scarce [11,34,35]. In order to compare the properties of the redox-active centers of pro- and eukaryotic XOs/XDHs, ¢rst EPR studies on XDH from P. putida 86 were performed, and the oxidation^reduction potentials of its redox-active centers were determined. 2. Materials and methods 2.1. Materials Bio-Scale DEAE-10 Macro-Prep MP10 and BioPrep SE-1000/17 were from Bio-Rad, Mu«nchen, Germany. Phenyl-Sepharose CL-4B was obtained from Pharmacia Biotech, Freiburg, Germany. Vivaspin 4 and Vivaspin 15 concentrators (molecular weight cut-o¡ (MWCO) 50 000) for ultra-¢ltration were from Vivascience Ltd., Binbrook Lincoln, UK. Precoated silica thin layer chromatography (TLC) sheets Polygram SIL G/UV254 were obtained from Macherey-Nagel, Du«ren, Germany. High-range and mid-range protein MW markers for sodium dodecyl sulfate^polyacrylamide gel electrophoresis (SDS^ PAGE) were purchased from Promega Corporation, Madison, WI, USA. All chemicals were of analytical grade. 2.2. Organism and growth conditions P. putida 86 [12] was grown in Erlenmeyer £asks in

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Table 1 Some properties of prokaryotic xanthine oxidoreductases and of two eukaryotic XOs Enzyme

Source

Molecular mass (kDa)

Subunit molecular mass(es) (kDa)

Subunit structure

Pyranpterin molybdenum cofactor

FeS cluster(s)

XDH

P. putida 86

550

91.0, 46.2

K4 L4

Mo-MPTa ((3.9) Mo)

XDH

P. synxantha 540 A3 C. acidovorans 287

76, 54

n.d.

90, 60

n.d.d (K4 L4 ?) K2 L2

R. capsulatus B10S R. capsulatus AI P. putida Fu1

275

83.2, 49.3

K2 L2

(2) Mo-MPT ((2.2) Mo) Mo-MPT

FeSI, FeSIIb (4.1) FAD (14.5) Fe, (15.8) Sc n.d. n.d. (`iron-containing £avoprotein') FeSI, FeSII (2) FAD (8) Fe FeSI, FeSII FAD

345

84.2

K4

(1.4) Mo

300

55, 25

K4 L4

(1.6) Mo

Ferricyanidelinked XDH XDH

P. putida 40

255

72

V. atypica

129

K3 (or K4 ) 82.4, 28.5, 18.4 KLQ

XDH

E. barkeri

530

81, 30, 17.5

K4 L4 Q4

XDH

S. cyanogenus

125

67

K2

XO

Arthrobacter sp. S-2 cow's milk

146

79

K2

V300

V150

K2

(Mo) (n.d.) (2) Mo-MPT

146.9

K2

(2) Mo-MPT

XDH XDH XDH XDH

XO XO

D. melanogaster 300

(Mo) (n.d.) Mo-MCDe ((0.86) Mo) MPT dinucleotide ((2.3) Mo; (1.1) W) ( 6 0.41) Mo

(8.4) Fe, (10.5) S (5.8) Fe, (2.4) S ^ (n.d.) (1.75) Fe, (1.61) S (17.5) Fe, (18.4) S (0.95) Se (14.3) Fe, (1.55) S (Fe, S) (n.d.) FeSI, FeSII (8 Fe), (8) S FeSI, FeSII

Flavin, or other cofactor

Reference

[13], this work [43] [35] [45]

(1.8^2) FAD

[44]

(0.9) Cyt. b

[9]

heme

[48]

(0.68) FAD

[21]

(2.8) FAD

[22]

(1.65) FAD

[24]

(FAD) (n.d.) (2) FAD

[49] [1,7,46]

(2) FAD

[63]

31

Mol mol of enzyme in parentheses, if reported. a Mo-MPT: molybdenum-molybdopterin. b FeSI: [2Fe2S] center showing smaller anisotropy in g tensors; FeSII: [2Fe2S] center showing larger anisotropy in g tensors. c S: acid-labile sul¢de. d n.d.: not determined. e Mo-MCD: molybdenum-molybdopterin cytosine dinucleotide.

mineral salts medium [15] containing 22 mM hypoxanthine as sole source of carbon, nitrogen, and energy. The cultures, which were incubated on an orbital shaker (140 rpm) at 30³C, were fed twice with hypoxanthine (after 14 and 22 h). Biomass was harvested after about 37 h of cultivation by centrifugation at 4400Ug, and stored at 380³C. To determine the level of constitutive XDH synthesis, P. putida 86 was grown in high nutrient broth (per liter: 5 g peptone, 5 g NaCl, 5 g yeast extract, 5 g meat extract).

2.3. Preparation of cell extract and puri¢cation of XDH Cell extracts of P. putida 86 were prepared by ultrasonic treatment and subsequent centrifugation as described previously [13]. For puri¢cation of XDH, the method described by Hettrich and Lingens [13] was modi¢ed. Crude extract obtained from 13 g cells (wet biomass) was fractionated by ammonium sulfate precipitation (0.9^1.8 M (NH4 )2 SO4 ). The precipitate was solubilized in 50 mM 2-amino-2-(hydroxymeth-

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yl)-1,3-propanediol (Tris)^HCl, pH 7.8, containing 0.6 M (NH4 )2 SO4 , and applied to a column of phenyl-Sepharose CL-4B (1.25U7 cm) that had been equilibrated in 50 mM Tris^HCl, pH 7.8, 1 M (NH4 )2 SO4 . The column was washed with 30 ml of 50 mM Tris^HCl, pH 7.8, containing 0.6 M (NH4 )2 SO4 , and with 70 ml of Tris^HCl bu¡er containing 0.4 M (NH4 )2 SO4 . XDH was eluted at a £ow rate of 1 ml min31 with a linear gradient of 0.4 M (NH4 )2 SO4 in 50 mM Tris^HCl pH 7.8 to 50 mM Tris^HCl pH 9.0 supplemented with 40% (v/v) glycerol (gradient volume: 180 ml), followed by a washing step with the latter, glycerol-containing bu¡er. Ammonium sulfate precipitation and hydrophobic interaction chromatography were performed at 4³C, whereas the subsequent chromatographic steps were carried out at room temperature. Fractions from the phenyl-Sepharose containing XDH activity were washed with 50 mM Tris^HCl, pH 7.4, and applied to a DEAE Macro-Prep MP10 column equilibrated in the same bu¡er. The column was washed with 20 ml of equilibration bu¡er followed by 20 ml of 50 mM Tris^HCl, pH 7.4, containing 0.2 M NaCl. XDH was desorbed in a linear gradient of 0.2 M NaCl to 0.8 M NaCl in 50 mM Tris^HCl, pH 7.4 (gradient volume: 200 ml). Active fractions were washed with 50 mM sodium phosphate bu¡er (pH 7.0) supplemented with 150 mM NaCl, and applied to a Bio-Prep SE-1000/7 column. Gel ¢ltration was performed at a £ow rate of 0.5 ml min31 in the same bu¡er. Fractions containing XDH were concentrated by ultra-¢ltration (4³C), and glycerol was added to a ¢nal concentration of 10% (v/v) prior to storage at 380³C. 2.4. Protein determination Protein concentrations were estimated by the method of Lowry et al. [36] using bovine serum albumin as standard protein. 2.5. Enzyme assays The reaction mixture of the standard assay (1 ml) contained in 50 mM Tris^HCl pH 8.0, 0.5% Triton X-100: 1 mM 2-(p-iodophenyl)-3-(p-nitrophenyl)-5phenyl-2H-tetrazolium chloride (INT), 0.03 mM Meldola's blue, and 1 mM xanthine. The reaction

was started by addition of enzyme preparation. The reduction of INT was followed at 503 nm. One unit (1 U) was de¢ned as the amount of enzyme that reduces 1 Wmol of INT per min at 25³C (INT^formazan, O503 nm = 19.3U103 M31 cm31 [37]). For the detection of XDH activity towards NAD‡ , the arti¢cial electron acceptor INT and the mediator Meldola's blue were replaced by 1 mM NAD‡ , and Triton X-100 was omitted from the test bu¡er. One unit (1 U) was de¢ned as the amount of enzyme that reduces 1 Wmol of NAD‡ per min at 25³C (NADH, O340 nm = 6.3U103 M31 cm31 [38]). To determine XO activity, urate formation was measured spectrophotometrically at 295 nm (O295 nm = 9.6U103 M31 cm31 [39]). The assay mixture contained bu¡er, 1 mM xanthine, and suitable amounts of enzyme. To determine which arti¢cial electron acceptors are used by XDH, the INT^Meldola's blue mix in the standard assay was replaced by INT, Meldola's blue (O569 nm = 13.9U103 M31 cm31 ), ferricyanide (O420 nm = 1.02U103 M31 cm31 [40]), phenazine methosulfate (O388 nm = 22U103 M31 cm31 [40]), 3,3P-(3,3P-dimethoxy-4,4P-biphenylylene)-bis[2-(p-nitrophenyl)-5-phenyl-2H-tetrazolium chloride] (NBT) (O566 nm = 15.5U103 M31 cm31 ), methylene blue (O610 nm = 41U103 M31 cm31 [40]), 2,6-dichlorophenol indophenol (O600 nm = 21U103 M31 cm31 [40]) (1 mM each), or 0.02 mM cytochrome c (O550 nm = 21U103 M31 cm31 [41]). 2.6. Determination of substrate speci¢city and identi¢cation of formed products The activity of XDH towards N-heteroaromatic compounds and aldehydes was determined spectrophotometrically in the standard assay. To identify the products of the XDH-catalyzed reactions, Meldola's blue and Triton X-100 were omitted from the standard assay, and 2 mM of putative substrate was added instead of 1 mM xanthine. Each assay contained 0.114 U of electrophoretically pure XDH and was incubated at 25³C until the INT^formazan had precipitated. After a brief centrifugation step, the supernatant was acidi¢ed and extracted with ethylacetate. The extract was concentrated in a rotary evaporator and analyzed by TLC using toluene/ dioxane/acetic acid, 72/16/1.6 (v/v/v) as mobile phase.

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2.7. PAGE PAGE under non-denaturing conditions was performed as described by Hames [42], using the highpH discontinuous bu¡er systems with 7.5% polyacrylamide in the resolving gel. For activity staining of XDH, gel slices were immersed in standard assay mixture lacking Meldola's blue. SDS^PAGE in 7.5%, 10%, or 12.5% polyacrylamide gels was performed according to Hames [42]. Gels were stained in 0.1% (w/v) Coomassie blue G in 50% (w/v) trichloroacetic acid, and de-stained in methanol/acetic acid/water, 30/10/60 (v/v/v). 2.8. Molecular mass estimation Matrix-assisted laser desorption/ionization-time of £ight-mass spectrometry (MALDI-TOF-MS) was performed with a Bruker Re£ex1 spectrometer. The matrix consisted of 3,5-dimethoxy-4-hydroxycinnamonic acid, tri£uoroacetic acid, and acetonitrile. Laser pulses (337 nm) were applied in 3 ns intervals at an intensity of 107 ^108 W cm32 . The molecular mass of the denatured enzyme was also determined by SDS^PAGE. 2.9. Preparation of EPR samples and EPR spectroscopy The samples for X-band measurements (200 Wl) were ¢lled into EPR quartz tubes (Wilmad) and immediately frozen in liquid nitrogen within 1 min, for Q-band samples (40 Wl) the ¢lling and freezing time was 3^4 min. XDH `as isolated' (5 mg ml31 ) was measured at 77 K. XDH was reduced by addition of an excess of sodium dithionite, or by addition of 500 nmol of xanthine to 1 nmol of enzyme under aerobic conditions. These samples were measured at 77 K, 20 K and 15 K. XDH (1 nmol) was also reduced under anaerobic conditions by addition of xanthine (500 nmol), frozen anaerobically, and measured at 70 K and 15 K. EPR spectra at X-band frequencies (9.5 GHz) were recorded on a Bruker ESP300 spectrometer equipped with a quartz dewar for measurements at liquid nitrogen temperature (77 K), or with a continuous helium £ow cryostat (ESR 900, Oxford instruments) for measurements at 15 K. The magnetic ¢eld and the microwave frequency

155

were determined with a nuclear magnetic resonance gaussmeter and a microwave counter, respectively. The modulation amplitude for spectra recording generally was 0.5 mT. For measurements at 77 K, several spectra at di¡erent microwave powers (0.2^10 mW) were recorded to avoid saturation broadening of the FAD radical signal. The spectra at 15 K were recorded at about 10 mW microwave power. For Qband experiments at 34.4 GHz, a QT-resonator from Bruker was used in a CF935 helium cryostat operated at 20 K. The spectra recorded on the ER220 spectrometer (Bruker) were digitized by home-built hard- and software. The modulation amplitude was 0.25 mT at 100 kHz, the microwave power was 0.28 mW. To improve the signal/noise ratio, Xand Q-band spectra were accumulated up to 50 times. Absolute signal intensities were determined by double integration using Cu2‡ -EDTA as standard. For simulation of the FeS spectra, the program Win-EPR Simfonia, Version 1.25 from Bruker Instruments was used. All spectra shown were baseline-corrected. 2.10. Redox titrations Redox titrations of XDH were carried out in 0.2 M Tris^HCl bu¡er, pH 8, at room temperature under strictly anaerobic conditions. The concentration of XDH in the titration experiments was 4.9 mg ml31 . The titration vessel was installed in an anaerobic chamber (atmosphere of 95% N2 , 5% H2 ), and stirred under oxygen-free argon gas. The redox potential was measured by a combined micro platinum electrode (Metrohm 6.0408.100) with an Ag/ AgCl reference, calibrated against a saturated quinhydrone solution, pH 7.0. The redox potential of the system was adjusted with small additions of 0.01 or 0.1 M solutions of Na2 S2 O4 or K3 [Fe(CN)6 ]. After equilibration for about 10 min at a certain redox potential, a sample volume of 250 Wl was withdrawn from the reaction vessel, transferred into an EPR tube and frozen immediately in liquid nitrogen under anaerobic conditions. In order to obtain stable potentials, the mediators described in Canne et al. [16] were used. The spectra of the iron^sulfur centers were recorded at 15 K, those of FAD and Mo(V) rapid species at 77 K. For the latter two signal giving centers, care was taken to avoid spectral broadening.

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For the Mo(V) center, the midpoint potential was determined from the intensity of the axial lines because of overlap of the ge component with the FAD signal. The redox behavior of the FeSI center was determined by evaluating the intensity of the nearly axial g tensor components (g2;3 ). Due to the low sample concentration and the broad and weak signals of FeSII, its redox potentials remained inaccessible. The midpoint potentials were obtained by simulation of the titration curves as described recently [16]. 3. Results and discussion 3.1. Induction of XDH synthesis Crude extracts from cells of P. putida 86 grown in high nutrient broth contained 0.014 U of XDH per mg of protein, compared to a speci¢c XDH activity of 0.91 U mg31 in crude extracts from cells grown on hypoxanthine as sole carbon source, which corresponds to a 65-fold increase in activity. When crude extracts from high nutrient broth-grown cells and from hypoxanthine-grown cells were subjected to non-denaturing PAGE and gel slices were immersed in bu¡er containing INT and xanthine, in each crude extract a single protein band turned red due to formazan formation. Since the relative mobilities of these protein bands stained with INT plus xanthine were identical, the XDH expressed constitutively at low level presumably corresponds to the hypoxanthine-induced XDH.

3.2. Puri¢cation of XDH, and subunit molecular masses The results of the puri¢cation procedure are summarized in Table 2. The puri¢cation scheme previously reported by Hettrich and Lingens [13] with a recovery of 47% led to XDH that showed a speci¢c activity of 31 U mg31 (xanthine:INT/Meldola's blue assay). Using the modi¢ed puri¢cation procedure described above, the speci¢c activity of the XDH preparations was improved to 35 U mg31 (standard assay), however, the recovery was much lower (Table 2). When NAD‡ served as electron acceptor, the speci¢c activity of the XDH puri¢ed from P. putida 86 was 17.9 U mg31 in the xanthine:NAD‡ assay and 26.7 U mg31 in the hypoxanthine:NAD‡ assay. XDHs from Pseudomonas synxantha A3, R. capsulatus AI (formerly: Rhodopseudomonas capsulata AI) and R. capsulatus B10S were reported to show speci¢c activities in the hypoxanthine:NAD‡ assay of 20 U mg31 [43], 35.9 U mg31 [44], and 22.6 U mg31 [45], respectively. XDH from C. acidovorans exhibited a higher activity of 50 U mg31 (xanthine:NAD‡ assay) [35]. In contrast to the XDHs from these aerobic bacteria, the selenium-containing XDH from E. barkeri exhibits a very high speci¢c activity of 164 U mg31 in the xanthine: NADP‡ assay [22]. The ratio A280 nm /A450 nm has widely been used as a criterion for purity in molybdenum-containing hydroxylases, a value of 5.0 corresponding to the pure enzyme (as exempli¢ed with milk XO) [1,7,46]. The ratios A280 nm /A450 nm = 4.96 and A450 nm / A550 nm = 2.96 observed with XDH puri¢ed from P. putida 86 are characteristic features of molybdenum

Table 2 Puri¢cation of XDH from P. putida 86 Puri¢cation step

Protein amount (mg)

Activity (U)

Speci¢c activity (U mg31 )

Puri¢cation (-fold)

Yield (%)

Crude extract Ammonium sulfate precipitation Phenyl-Sepharose DEAE-10 Macro-Prep MP10 Bio-Prep SE-1000/17

549 132 32 7.2 2.2

500 351 322 152 77.4

0.9 2.7 10.1 21.1 35.2

1 3.0 11.2 23.5 39.1

100 70 64 30 15

Starting material was 13 g of wet biomass. XDH activity was determined in the standard assay with INT as electron acceptor and Meldola's blue as mediator.

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hydroxylases containing the typical cofactor stoichiometry of the eukaryotic XDHs/XOs (Table 1) and indicate the purity of the enzyme preparation. The XDH preparations were electrophoretically homogeneous. SDS^PAGE of XDH showed two protein bands with molecular masses of approximately 94 and 51 kDa. MALDI-TOF-mass spectra of XDH showed two prominent peaks at m/z = 91 041 and m/z = 46 238, corresponding to the molecular masses of the subunits. Since the molecular mass of native XDH had been estimated to 550 kDa, an K4 L4 structure seems plausible [13]. XDH from P. synxantha A3, which consists of 54 kDa and 76 kDa subunits, may also form an K4 L4 structure [43]. The XDHs form C. acidovorans and from R. capsulatus B10S likewise possess two subunits of roughly comparable size, but these obviously form an K2 L2 native structure [35,45]. The molecular mass of each KL protomer of these prokaryotic XDHs corresponds to the monomeric molecular mass (about 150 kDa) of the eukaryotic enzymes (Table 1). The 46 kDa subunit of XDH from P. putida 86 may represent a fusion of the small and the medium-sized subunit of the KLQ- or (KLQ)2 -type bacterial molybdenum hydroxylases. The KLQ composition has so far been found only for the XDH from V. atypica [21], however, XDH from E. barkeri is also composed of three di¡erent subunits of comparable sizes, but these appear to form an K4 L4 Q4 structure [22] (Table 1). 3.3. Speci¢city towards N-heteroaromatic compounds and aldehydes The relative activities of XDH from P. putida 86

toward some N-heteroaromatic compounds and aromatic aldehydes are shown in Table 3. Some of the N-heteroaromatic compounds tested were regiospeci¢cally hydroxylated at the electropositive carbon atom adjacent to the N-heteroatom. However, quinoline, isoquinoline, quinaldine, nicotinate, 6-chloronicotinate, imidazole, uracil, pyridine and pyrimidine were not converted by XDH. Acetaldehyde, butyraldehyde, and 4-hydroxy-3-methoxybenzaldehyde (vanillin) likewise were not oxidized by XDH. Similar to the XDH from P. putida 86 (Table 3), XDHs from P. synxantha A3 [43], Streptomyces cyanogenus [24], and R. capsulatus AI [44] show highest activity toward hypoxanthine (133%, 164%, and 149% relative activity, respectively, compared with the activities toward xanthine, 100%). In contrast, XDHs from C. acidovorans [47], P. putida Fu1 [9], P. putida 40 (ferricyanide-linked [48]), and Arthrobacter sp. S-2 (ferricyanide assay [49]) were more active toward xanthine than toward hypoxanthine. Besides their relatively high activity toward hypoxanthine, another common feature of the XDHs from P. putida 86, P. synxantha A3 [43], and S. cyanogenus [24] is their low activity toward purine as reducing substrate. The relative activities of milk XO with xanthine, hypoxanthine, and purine were 100%, 76%, and 59%, respectively (ferricyanide assay [23]). XDH from P. putida 86 slowly converted benzaldehyde and some substituted benzaldehydes to the corresponding benzoates (Table 3), whereas the aliphatic aldehydes tested did not serve as substrate. Aromatic aldehydes are also oxidized by the XDHs from V. atypica (benzaldehyde: 4%, salicylaldehyde: 2% relative activity [21]) and S. cyanogenus (benzal-

Table 3 Speci¢city of XDH from P. putida 86 towards reducing substrates Substrate

Relative activity (%)

Product formed

Xanthine Hypoxanthine Purine Quinazoline Phthalazine Benzaldehyde 3,4-Dihydroxybenzaldehyde 4-Hydroxybenzaldehyde 4-Dimethylaminobenzaldehyde

100 149 5.0 2.0 0.4 1.3 1.2 0.7 0.3

urate urate n.d.a 4-(3H)quinazolinone 1-(2H)phthalazinone benzoate 3,4-dihydroxybenzoate 4-hydroxybenzoate 4-dimethylaminobenzoate

a

157

n.d.: not determined.

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dehyde: 0.16% relative activity [24]) (all values are relative to the activity toward xanthine). However, the latter two enzymes are even more active toward short-chain aliphatic aldehydes: acetaldehyde is oxidized by XDHs from V. atypica and S. cyanogenus with relative activities of 5% [21] and 1.2% [24], respectively (the activity toward xanthine set as 100%). XO from cow's milk is capable of oxidizing a wide variety of aldehydes. Simple aliphatic aldehydes are converted rather rapidly, acetaldehyde, which is oxidized with a relative activity of 25% as compared with the activity toward xanthine, being the best substrate (ferricyanide assay [23]). The broad speci¢city toward reducing substrates of some molybdenum hydroxylases, especially of bovine XO, led to the idea that these enzymes may have more general functions in vivo, such as detoxi¢cation of cytotoxic compounds, and/or oxidation of various N-heteroaromatics (substituted purines and pyrimidines) as potential nutrients. However, the physiological role of the observed di¡erences in substrate speci¢cities of XDHs/XOs from di¡erent organisms remains uncertain. The broad substrate speci¢city suggests that there are few steric constraints in the vicinity of the molybdenum center. However, since site-directed mutagenesis studies on molybdenum hydroxylases that attempt to identify amino acid residues responsible for binding and positioning of the reducing substrate are scarce [50], the molecular basis of substrate speci¢city is not understood. 3.4. Electron acceptor speci¢city The e¤ciencies of several electron acceptors in facilitating the oxidation of xanthine by XDH from P. putida 86 are shown in Table 4. Of the oxidizing substrates tested, NAD‡ is the most e¤cient, followed by INT and ferricyanide. The oxidase activity of the enzyme was 25.6-fold lower than its dehydrogenase activity (Table 4). In rat liver XDH, the conversion of dehydrogenase to oxidase results from an altered conformation of the enzyme, and involves a change in the oxidation^reduction behavior of the £avin. The dehydrogenase form of the enzyme maintains a stable £avin semiquinone signal when reduced by xanthine, whereas in the oxidase form, the fully reduced £avin is formed preferentially. In the fully reduced level, the rat liver enzyme is thought to lose

Table 4 Speci¢c activity of XDH from P. putida 86 with oxidizing substrates Redox pair

E0 P

[Fe(CN)6 ]33 /[Fe(CN)6 ]43 +360a O2 /H2 O2 +295a Cytochrome c, ox/red +256b 2,6-Dichlorophenol indophenol, ox/red +217a INT, ox/red +90c Phenazine methosulfate, ox/red +80a NBT, ox/red +50c Methylene blue, ox/red +11a Meldola's blue, ox/red 38d ‡ NAD /NADH 3320a

Speci¢c activity (U mg31 ) 10.4 0.7 0.1 b.d.e 16.4 0.1 1.0 b.d.e 2.1 17.9

a

[70]. [71]. c [72]. d [73]. e b.d.: activity was below the limit of detection. b

its ability to interact with NAD‡ and to acquire the enhanced capacity to transfer electrons to O2 [51]. The observed low rate of cytochrome c reduction in the presence of XDH and xanthine may be due to traces of superoxide anion that may be formed in the oxidase reaction [7,52,53]. Using 2,6-dichlorophenol indophenol as possible electron acceptor, the UV/Vis absorption spectrum of the assay mixture changed upon addition of enzyme, but urate formation was not observed. 2,6-Dichlorophenol indophenol, which is thought to interact with the molybdenum center [54], has been reported to act as a competitive inhibitor for milk XO [55,56]. Whereas O2 and NAD‡ are known to react at the £avin site of XOs and XDHs [7,57,58], electron transfer to phenazine methosulfate [7,59] and to ferricyanide [7,57] is presumed to take place from an iron^sulfur center. However, the speci¢c activity with ferricyanide in the aerobic assay may additionally involve the non-enzymatic reduction of ferricyanide by the reaction product urate [60]. NBT is also thought to react at an iron^sulfur cluster, but additionally it might be reduced by superoxide anion radical [7,61]. XDH from P. synxantha A3, which resembles XDH from P. putida 86 with respect to its high molecular mass and its activity toward the reducing substrates xanthine, hypoxanthine, and purine, di¡ers

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from the P. putida 86 enzyme with respect to electron acceptor speci¢city, since the enzyme from P. synxantha A3 is unable to utilize ferricyanide, INT, and dioxygen as oxidizing substrates [43]. Speci¢city of vertebrate XDHs with regard to electron acceptors is very broad and shows di¡erent speci¢city patterns [7]. The redox potentials of the £avin chromophores in the oxidases and dehydrogenases determine whether dioxygen or NAD‡ is used e¤ciently as an oxidizing substrate (see Section 3.6 and Tables 4 and 5). For the XDH from P. putida 86, NAD‡ seems to be the physiological electron acceptor. 3.5. EPR spectroscopy XDH from P. putida 86 in the oxidized state (as isolated, aerobic conditions) did not elicit any EPR signal (Fig. 1a). Upon reduction of aerobically prepared samples by addition of an excess of dithionite, a rather weak signal of the FAD semiquinone radical (g = 2.0037) was observed at 77 K showing a line width of 1.45 mT (Fig. 1b). Surprisingly, after addition of the substrate xanthine, a signal with a line width of 2.1 mT was measured for the FAD signal (Fig. 1c). These two values are characteristic for the red (anionic, 1.5 mT) and the blue (neutral, 1.9 mT) form of the FAD radical, respectively [62]. Compared to the UV/Vis absorption spectrum of XDH as isolated (oxidized form), the spectrum of the xanthine-reduced enzyme showed an increase in absorption at 620 nm along with a decrease in absorption at 450 nm, which also indicates the neutral £avin semiquinone. It appears that in the case of xanthine, a protonation of the FAD moiety is occurring whereas in the presence of dithionite the deprotonated state is prevailing. The origin for the di¡erent behavior upon reduction with xanthine and dithionite is presently not clear. In addition, there are also small features at higher ¢elds visible, which can be associated to the Mo(V) rapid type signal because of the characteristic splitting of 1.4 mT and the corresponding perpendicular g factor of 1.965 typically also found in other molybdenum hydroxylases like quinoline 2-oxidoreductase from the same P. putida strain 86 [15,16], turkey or chicken liver XDH [28,29], Drosophila melanogaster XDH [63], or milk XO [1]. The other principal component (ge ) is not resolved in the £ank of the FAD signal.

159

When the temperature is decreased to 15 K, resonances from [2Fe2S] centers become more pronounced (Fig. 2). One expects spectra from two different [2Fe2S] centers, which, according to their anisotropy in g tensors, are designated FeSI (for smaller anisotropy) and FeSII (for larger anisotropy). The experimental patterns are, however, dominated by the rather broad signal in the region around g = 2, where usually the FAD radical is found (Fig. 2a^d) and thus make a di¡erentiation di¤cult, especially for FeSI. It is noted that line broadening e¡ects are occurring on the FAD signals under the conditions of low temperatures and relatively high microwave powers (10 mW) necessary to observe the FeS species. In the three experimental spectra obtained at

Fig. 1. X-band EPR spectra of XDH as isolated (a), and reduced under aerobic conditions with dithionite (b) and the substrate xanthine (c) were recorded at 77 K. The dominant line at g = 2.0037 arises from the FAD radical species. Small contributions of a Mo(V) rapid-like signal are marked with arrows. The additional short traces show the Mo(V) features with lower noise.

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Fig. 2. The X-band EPR spectra of XDH reduced under aerobic conditions with dithionite (a) and xanthine (b) and under anaerobic conditions with substrate (d) were obtained at 15 K. At this temperature, both FeS signals become visible. Spectrum (c) represents a simulation of spectrum (b) with a rhombic FeSII species (2.085, 1.99, 1.907) and a nearly axial FeSI species (2.016, 1.926, 1.924) reproducing the resonances of the FeS signals. The stick diagrams indicate the position of the g components. The g2 component of FeSII and the g1 resonance of FeSI are overlapping with the FAD signal. Spectrum (e) was taken at 70 K showing the FAD radical signal and small contribution of Mo(V) rapid-like signal.

X-band frequencies and 15 K (Fig. 2a,b,d), FeSII shows an extremely broad asymmetric resonance at g1 W2.085 with a line width of ca. 5.5 mT and a g3 component at 1.905. Its g2 component is not resolved but may be buried under the dominant line (with gW1.99). Very similar g factors for the FeSII center are found in other molybdenum hydroxylases, like quinoline 2-oxidoreductase from the same P. putida strain 86 [15], eukaryotic XOs/XDHs [1,28,29,63], Aspergillus nidulans purine hydroxylase II [30], and carbon monoxide dehydrogenase from carboxydo-

trophic bacteria [64]. In the spectra, an additional biphasic signal (above and below the zero line, marked with asterisks in traces a and b) is present, which either corresponds to the g2 component of a rhombic spectrum or to an axial component. In order to distinguish the two possibilities, spectral simulations were performed with either two rhombic or one rhombic and one axial FeS species. The spectral features of this line (and of the weak high ¢eld resonance) are best reproduced by assuming a nearly axial signal (g2 = 1.929, g3 = 1.924) arising from the FeSI species as shown in the simulation of trace c in Fig. 2. Its other component (g1 W2.016) is located within the dominant line superimposing with the signals of FAD and a presumably broad g2 component of FeSII. These g factors are comparable to those of the axial FeSI center described for quinaldine 4-oxidase (ge = 2.021, gP = 1.937) [16] and represent a second example of a FeSI species with axial symmetry. (However, FeSI of A. nidulans purine hydroxylase II has also been reported to be of near axial symmetry [30]). Raising the temperature of the anaerobically prepared XDH reduced with substrate from 15 K to 70 K leads to a fading of the FeS signals, and, apart from an intense FAD signal, the axial features of a Mo(V) rapid signal were clearly observable at gP = 1.965, AP = 1.4 mT (Fig. 2e) as already found for the other samples at 77 K. Again the ge feature of the rapid signal is barely resolved in the £ank of the FAD signal. The relative intensity of the Mo(V) signal was slightly higher in the anaerobic sample than in the aerobic XDH preparations. There are some indications of the superposition of spectral species in the rather broad intense line around g = 2 as can be inferred from the shoulders and line distortions in spectra a and d of the dithionite- and xanthine (anaerobically)-reduced samples (Fig. 2). In order to increase the spectral resolution, Q-band measurements were performed at 20 K (Fig. 3). For the dithionite-reduced sample (trace a) again the intense feature arising from the FAD signal is observed. The axial part of the FeSI signal exhibits a small rhombic contribution which is simulated in spectrum b (with g2 = 1.930, g3 = 1.924). From a comparison with spectrum c (xanthine-reduced), it is evident that its signal intensity is diminished. Concomitantly, the broad shoulder low ¢eld of FAD is also reduced indicating the position of the g1 com-

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ponent of FeSI, which is simulated in spectrum b with g1 = 2.024. The spectral features of FeSI are aligned via the dotted lines. In X-band spectra (Fig. 2), this signal remains unresolved under the broad resonance around g = 2. For the FeSII center, the peculiar line shape observed in X-band is also found in Q-band spectra. The g1 feature shows a large width of nearly 40 mT with an asymmetric low ¢eld slope and indications of additional species present (Fig. 3a,c). Moreover, an additional resonance is resolved in spectrum c and corresponds to a shoulder-like feature in trace a (marked with an arrow in Fig. 3a,c). It may either represent the g2 component of FeSII or an other species. In the ¢rst case, the g factor would be exceptionally high (2.04) for this type of center which has not been observed in any other related enzyme. Similar to the X-band spectra, the g3 resonance can be seen (with low intensity in spectrum c) in the Q-band experiments. However, also from Q-band spectra, a clear assignment of the g2 component of FeSII cannot be derived. The apparent line width for the g = 2 radical signal in the Q-band spectra amounts to 3.5 and 4 mT and is smaller than the line width of 4.5 mT found in the X-band spectra. This is related to the reduced overlap with iron^sulfur center signals and to a less pronounced spectral broadening. On the whole, the EPR-active centers in XDH exhibit a high similarity to other enzymes of the XO family. The FeS centers are observable only at temperatures below 60 K. FeSII shows typical spectral properties relating to g factors and the observed large line widths. The spectral shape observed in Qband spectra indicates that FeSII is constituted of several still overlapping species of various intensities. A distinct assignment of their principal g factors is therefore not achievable. With respect to FeSI, the enzyme appears to be close to quinaldine 4-oxidase [16]. With manual freeze preparation ( s 30 s), the only Mo species present in rather low amounts is a rapid-like form, which has been assigned to Mo(V) with no substrate bound [16,32]. No traces of the kinetically important intermediate Mo(V) very rapid species were detectable, which resembles the ¢ndings for milk XO [1,32], turkey liver XDH [28], or bacterial quinoline 2-oxidoreductase and quinaldine 4-oxidase [18], where the very rapid species was clearly resolvable only by rapid freeze methods.

161

Fig. 3. The Q-band (34.4 GHz) EPR spectra of XDH reduced under aerobic conditions with dithionite (a) and xanthine (c) were measured at 20 K. The asterisks in spectrum a indicate positions of Mn2‡ lines which are observed from the Q-band cavity at low temperatures and long accumulation times. Spectrum b represents the simulation of the FeSI center related to the experimental patterns by the dotted lines. The stick diagram marks the region of the g1 components and g3 of FeSII. The resonance of unclear origin is indicated by arrows.

3.6. Oxidation^reduction potentials of the redox-active centers Redox titrations of XDH were carried out in TrisHCl bu¡er at pH 8. It was not possible to determine the redox potential of the FeSII center, since g1 and g3 were too weak to allow quantitative determination, and g2 W1.99 severely overlapped with mediator and FAD signals. Due to the relatively low amount of enzyme available and the high dilutions that were necessary, the magnitude of the errors associated

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Table 5 Oxidation^reduction potentials of redox-active centers of some XOs/XDHs Enzyme

XDH XDH XDH XDH XO XO XO XO XO XO

Source

P. putida 86 turkey liver chicken liver cow's milk cow's milk cow's milk cow's milk cow's milk cow's milk cow's milk

Bu¡er, pH

Tris, 8.0 pyrophos.b , 8.2 pot. phos.c , 7.8 sod. phos.d , 7.5 sod. phos., 7.5 bicinee , 7.7 bicine, 7.7 pot. phos., 7.8 Tris, pH 8.2 pyrophos., 8.2

Potential versus standard hydrogen electrode (mV)

Reference

Mo (VI/V)

Mo (V/IV)

FAD/ FADH

FADH / FADH2

FeSI ox/red

FeSII ox/red

3360 3350 3357 n.d. n.d. 3345 3373 3355 3397 3355

3300 3362 3337 n.d. n.d. 3315 3377 3335 3405 3355

3350 3359 3345 3270 3280 3301 3332 3310 3378 3351

3350 3366 3377 3410 3230 3237 3234 3220 3223 3236

3335 3295 3280 3310 3310 3310 3310 3280 3336 3343

n.d.a 3292 3275 3235 3235 3217 3255 3245 3255 3303

this work [74] [29,75] [65] [65] [68] [67] [75] [69] [69]

a

n.d.: not determined. pyrophos.: pyrophosphate bu¡er. c pot. phos.: potassium phosphate bu¡er. d sod. phos.: sodium phosphate bu¡er. e bicine: N,N-bis(2-hydroxyethyl)glycine. b

with the measured potentials were of the order of þ 30 mV, except for the FeSI potential, which is more accurate ( þ 15 mV). The oxidation^reduction potentials determined for the various centers in the P. putida 86 XDH resemble those of the corresponding centers in eukaryotic XDHs (Table 5), suggesting that the environment of each center and their functionality in these enzymes are quite similar. The midpoint potentials determined for the molybdenum, FAD and FeSI redox couples of the bacterial XDH are close to each other. The £avin midpoint potential of 3350 mV is su¤ciently low for reduction of NAD‡ (with a midpoint potential of the NAD‡ /NADH couple of 3335 mV at pH 7.5 [65]). A detailed mechanism for electron distribution within the enzyme (`rapid equilibrium hypothesis') based on the relative potentials of the redox-active centers has been proposed by Olson et al. [66] for milk XO. It is noted that the di¡erences in the reported values (Table 5) may, in part, be caused by pH e¡ects, since variations of the midpoint potentials with changes in pH have been observed for all redox-active centers of XO [29,67,68]. There is also an e¡ect of the bu¡er system used and a temperature e¡ect in the measured potential [68,69].

4. Conclusions XDH from P. putida 86 catalyzes the oxidation of hypoxanthine, xanthine, purine, and some aromatic aldehydes, using NAD‡ as the preferred electron acceptor. It resembles XDH from P. synxantha A3 with respect to its molecular mass, presumed subunit composition, and speci¢city toward some reducing substrates, but the speci¢cities toward oxidizing substrates di¡er considerably in these two bacterial XDHs. Generally, XDHs/XOs from prokaryotic sources are very diverse with respect to their molecular masses, subunit structures, cofactor compositions, and substrate and electron acceptor speci¢cities. The enzyme from P. putida 86 has the same cofactor composition as eukaryotic XDHs/XOs, containing Mo-MPT, two distinct [2Fe2S] clusters, and FAD. Both the oxidation^reduction potentials and the EPR characteristics of the redox-active centers of P. putida 86 XDH are highly similar to the features observed in eukaryotic XDHs. An exception is the FeSI center, which in XDH from P. putida 86 shows nearly axial symmetry, whereas milk XO and most other molybdenum hydroxylases have a rhombic FeSI species.

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Acknowledgements We thank Dr. H.-J. Hecht, GBF Braunschweig, for MALDI-TOF-MS analyses. The ¢nancial support of the Deutsche Forschungsgemeinschaft and the Volkswagen-Stiftung is gratefully acknowledged.

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