Insights into the mechanism of dihydropyrimidine dehydrogenase from site-directed mutagenesis targeting the active site loop and redox cofactor coordination

Biochimica et Biophysica Acta 1804 (2010) 2198–2206

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Biochimica et Biophysica Acta j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / b b a p a p

Insights into the mechanism of dihydropyrimidine dehydrogenase from site-directed mutagenesis targeting the active site loop and redox cofactor coordination Bernhard Lohkamp a, Nina Voevodskaya b, Ylva Lindqvist a, Doreen Dobritzsch a,⁎ a b

Department of Medical Biochemistry and Biophysics, Karolinska Institutet, 171 77 Stockholm, Sweden Department of Biochemistry and Biophysics, Stockholm University, 106 91 Stockholm, Sweden

a r t i c l e

i n f o

Article history: Received 3 June 2010 Received in revised form 13 August 2010 Accepted 31 August 2010 Available online 8 September 2010 Keywords: Dihydropyrimidine dehydrogenase Flavoprotein Iron–sulfur clusters Site-directed mutagenesis Oxidoreductase Pyrimidine degradation

a b s t r a c t In mammals, the pyrimidines uracil and thymine are metabolised by a three-step reductive degradation pathway. Dihydropyrimidine dehydrogenase (DPD) catalyses its first and rate-limiting step, reducing uracil and thymine to the corresponding 5,6-dihydropyrimidines in an NADPH-dependent reaction. The enzyme is an adjunct target in cancer therapy since it rapidly breaks down the anti-cancer drug 5-fluorouracil and related compounds. Five residues located in functionally important regions were targeted in mutational studies to investigate their role in the catalytic mechanism of dihydropyrimidine dehydrogenase from pig. Pyrimidine binding to this enzyme is accompanied by active site loop closure that positions a catalytically crucial cysteine (C671) residue. Kinetic characterization of corresponding enzyme mutants revealed that the deprotonation of the loop residue H673 is required for active site closure, while S670 is important for substrate recognition. Investigations on selected residues involved in binding of the redox cofactors revealed that the first FeS cluster, with unusual coordination, cannot be reduced and displays no activity when Q156 is mutated to glutamate, and that R235 is crucial for FAD binding. © 2010 Elsevier B.V. All rights reserved.

1. Introduction Dihydropyrimidine dehydrogenase (DPD; EC 1.3.1.2) catalyses the first step in the reductive degradation of pyrimidines by reducing uracil and thymine in an NADPH-dependent reaction to 5,6-dihydrouracil and 5,6-dihydrothymine, respectively [1]. The dihydropyrimidines are subsequently further degraded to β-alanine and β-aminoisobutyric acid, respectively, under release of carbon dioxide and ammonia (Fig. 1a). In mammals the pathway is the major route for synthesis of β-alanine, a putative neurotransmitter and building block of the neuropeptides carnosine and anserine [2]. DPD also degrades 5-fluorouracil (5FU), a drug widely used in chemotherapy of e.g. breast, colorectal and head/neck cancer. The rapid breakdown of N80% of the administered 5FU necessitates high dosage [3]. In this context, a partial or complete DPD deficiency is a pharmacogenetic disorder affecting cancer patients who develop severe toxicity, including death, following the administration of 5FU [4,5]. Inhibitory drugs targeting human DPD are under development

Abbreviations: DPD, dihydropyrimidine dehydrogenase; DTT, dithiothreitol; WT, wildtype; EPR, electron paramagnetic resonance; GltS, glutamate synthase; PMSF, phenylmethylsulphonyl fluoride; PBS, phosphate buffered saline; HEPES, 4-(2-hydroxyethyl)-1piperazineethanesulfonic acid; Tris, tris(hydroxymethyl)aminomethane ⁎ Corresponding author. Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Tomtebodavägen 6, SE-171 77 Stockholm, Sweden. Tel.: +46 8 5248 7651; fax: +46 8 32 76 26. E-mail address: [email protected] (D. Dobritzsch). 1570-9639/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.bbapap.2010.08.014

or already utilized in anti-cancer therapy to increase 5FU efficiency and to allow administration of lower doses [6]. Liver DPD has been purified from natural sources [7–10] as well as recombinantly expressed [11]. The high sequence similarity between DPDs from different mammalian species (e.g. 93% between human and pig) suggests a very similar reaction mechanism and threedimensional structure. The highly modular structure of the recombinant pig liver DPD was revealed by X-ray crystallography [12]. Each subunit of the homodimeric enzyme (2× 110 kDa) harbours two flavins, FAD and FMN, as well as four [4Fe–4S] clusters (Fig. 2a and b). Electrons are transferred from a NADPH binding site in domain III (site 1) to the active site for pyrimidine reduction in domain IV (site 2) via a chain of FeS clusters. Based on spectroscopic, biochemical and structural data a nonclassical two-site ping-pong reaction mechanism could be deduced [13] (Fig. 1b). Upon NADPH binding a hydride is transferred from its C4-atom to atom N5 of FAD, which leaves NADP+ and FADH− at site 1. Transfer of two electrons to the FeS clusters is likely followed by proton release from atom N5 to complete the redox cycle of the FAD. The first two clusters in the chain, nFeS2 and nFeS1, are bound by the α-helical domain I of DPD (Fig. 2a, b and d), the other two, cFeS1 and cFeS2, are bound by domain V. Cluster nFeS2 is closest to the FAD and shows an unusual coordination by one glutamine and three cysteine residues (Fig. 2c). The FMN located in site 2 is reduced by transfer of electrons from cFeS2. The pyrimidine substrate is bound with the si-faces at atoms C6 and C5 directed towards the FMN-N5 and the catalytic cysteine C671, respectively. Transfer of the proton from C671 and hydride from FMN is proposed to occur in a concerted anti-addition reaction. C671 is located

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Fig. 1. (a) Reductive pyrimidine catabolic pathway. The reactions are catalysed by dihydropyrimidine dehydrogenase (DPD), dihydropyrimidinase (DHP, E.C. 3.5.2.2) and β-alanine synthase (β-AS, also called β-ureidopropionase, E.C. 3.5.1.6). (b) Scheme of the kinetic mechanism of DPD. First, NADPH binds to site 1 and reduces the FAD. This then transfers electrons via four FeS clusters to the FMN at site 2, which then reduces uracil to 5,6-dihydrouracil (DHU). Adapted from Schnackerz et al. [26].

in the so-called active site loop comprising residues 670–683, which has been observed in open and closed conformational states [12,14] (Fig. 2e). Only in the closed loop state C671 is appropriately positioned for proton transfer. To gain further insights into the catalytic mechanism of DPD we targeted five residues putatively important for enzyme function in site-directed mutagenesis experiments. S670 and H673 are located in the pyrimidine binding site and expected to be crucial for sensing its occupational state and for stabilizing the closed active site loop conformation. Q156 and C126 involved in ligation or near nFeS2 and the FAD-binding R235 were targeted to investigate their specific functions in redox cofactor binding, electron transfer and catalytic mechanism. 2. Material and methods 2.1. Materials The vector pET28 and E. coli strain Tuner(DE3) were purchased from Novagen, the vector pMOSBlue from GE Healthcare, and talon resin from Clontech. 2′,5′-ADP-Sepharose resin and Superdex S-200 column were obtained from Amersham Biosciences. All chemicals used were obtained from commercial sources and of the highest grade available.

with the same enzymes. For expression the vector pET28-pDPD was transformed into the E. coli strain Tuner(DE3). The desired mutations in the DPD gene were introduced using the QuikChange method (Stratagene). Here, PfuTurbo DNA polymerase (Stratagene) was used for amplification in the presence of 2% (v/v) DMSO and 8% (v/v) glycerol. The following primer pairs were used (mutation sites underlined): C126A primers, 5′-CCTCTTGGTCTGACCGCTGGAATGGTATGTCCAAC-3′ and 5′-GTTGGACATACCATTCCAGCGGTCAGACCAAGAGG-3′; Q156E primers, 5′-CAATTAATATTGGTGGATTGGAGCAGTTTGCTTCTGAGGTG-3′ and 5′-CACC TCAGAAGCAAACTGCTCCAATCCACCAATATTAATTG-3′; R235A primers, 5′-TCTGAAATCCCTCAGTTCGCGCTGCCATATGATGTAGTG-3′ and 5′-CACTACATCATATGGCAGCGCGAACTGAGGGATTTCAGA-3′; R235K primers, 5′-CTTCTGAAATCCCTCAGTTCAAGCTGCCATATGCTGTAGTGAA3′ and 5′-TTCACTACATCATATGGCAGCTTGAACTGAGGGATTTCAGAAG3′; S670A primers, 5′-CCTTGGAGTTAAATCTGGCATGTCCACACGGCATG3′ and 5′-CATGCCGTGTGGACATGCCAGATTTAACTCCAAGG-3′; H673N primers, 5′-TTAAATCTGTCATGTCCAAACGGCATGGGAGAAAGAG-3′ and 5′-CTCTTTCTCCCATGCCGTTTGGACATGACAGATTTAA-3′; H673Q primers, 5′-TAAATCTGTCATGTCCACAGGGCATGGGAGAAAGA-3′ and 5′-TCTT TCTCCCATGCCCTGTGGACATGACAGATTTA-3′. The sequence of the WT DPD gene and the site-specific mutations were confirmed by DNA sequencing. 2.3. Protein expression

2.2. Cloning and mutational analysis The gene for pig liver DPD was amplified by PCR using Platinum Pfx DNA polymerase (Invitrogen). The pSE420 vector containing pig DPD cDNA [11] was used as a template with the following primers: 5′-TAC TGCGATGCTAGCGCCCCTGTGCTGAGCAAGGACGTG-3′ and 5′-TACTGCGATCTCGAGTCAGcACACCGGATTCACAGCCAA-3′ (bold: NheI and XhoI restriction sites, respectively). The PCR fragment was cloned into a pMOSBlue vector. The DPD gene fragment was cut with the restriction enzymes NheI and XhoI, purified and ligated into a pET28 vector cut

Bacterial pre-cultures of E. coli Tuner(DE3) strain harbouring pET28pDPD were grown in LB medium in presence of 50 μg/ml kanamycin over night at 37 °C. 1 l auto-induction medium, supplemented with 80 μg/ml antibiotic and 100 μg/ml uracil, was inoculated with 10 ml of pre-culture and grown under agitation at 30 °C [15]. After 4 h of growth 10 μM sodium sulfide, 10 μM ammonium ferric citrate, 10 μM FAD and 10 μM FMN were added to the culture. Cells were harvested by centrifugation after further 20 h of growth and washed with PBS buffer. Cells were resuspended in 1 ml/g of cell lysis buffer (50 mM

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Fig. 2. Mutation sites in the structure of DPD. (a) Homo-dimer of DPD in ribbon representation with domains I, II, III, IV, and V coloured in green, cyan, gold, red, and blue respectively. Grey spheres indicate mutation sites and one subunit is shown transparent for clarity. (b) One of the two electron transfer chains of the DPD dimer. Subscripts A and B denote the parent protein chain of the compound. The orientation is the same as in (a). (c) Unusual FeS cluster coordination of nFeS2 in DPD. Solid lines indicate the cluster ligation. (d) NADPH binding in DPD. The green solid line indicates the hydride transfer, the red dotted line the proton shuttling path and black dotted lines the electron transfer path. (e) Stereo-view of the open (cyan; pdb code: 1h7x) and closed (green; pdb code: 1gth) active site loop conformation of DPD. Hydrogen bonds of S670 and H673 in the open and closed form are indicated by grey and black dotted lines, respectively.

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sodium phosphate pH 7.5, 300 mM NaCl, 5 mM imidazole, 5 mM β-mercaptoethanol, 0.1 mM PMSF, 1 mM benzamidine, 1 protease inhibitor tablet per litre) supplemented with 300 μg lysozyme/g of cells and 1 μg DNaseI/g of cells, flash frozen in liquid nitrogen and stored at −20 °C until further use. The yield is typically 20 g of wet cell weight per litre of culture medium. 2.4. Protein purification Resuspended cells were twice thawed and flash frozen in liquid nitrogen; more DNaseI was added if necessary to reduce the viscosity of the sample. The cells were then further disrupted by sonication at 4 °C. The homogenate was centrifuged at 30,000 g for 60 min. The supernatant was filtered with a 0.22 μm filter and the cleared lysate applied onto a Talon resin column (1.5 ml) by gravity flow. The pellet was resuspended in lysis buffer, sonicated, centrifuged and applied to the Talon column as described earlier. The column was connected to an FPLC system operating at a flow rate of 1 ml/min. The column was washed extensively with lysis buffer (approximately 30 column volumes) and elution was accomplished by applying a linear gradient over 30 column volumes of elution buffer (50 mM sodium phosphate pH 7.5, 300 mM NaCl, 150 mM imidazole, 5 mM β-mercaptoethanol, 0.1 mM PMSF, 1 mM benzamidine, 1 protease inhibitor tablet per litre). Fractions containing DPD were pooled and applied onto a 2′,5′ADP-Sepharose column (18 ml) equilibrated with low salt buffer A (35 mM Tris pH 7.5, 1 mM DTT, 0.1 mM PMSF, 1 mM benzamidine, 0.4 M KCl, 1 protease inhibitor tablet/litre). The column was washed with 3 column volumes of buffer A and elution was accomplished with a linear gradient over 10 column volumes to high salt buffer B (35 mM Tris pH 7.5, 1 mM DTT, 0.1 mM PMSF, 1 mM benzamidine, 2 M KCl). Mutant R235A was purified by gel filtration using a Superdex S-200 column (1.6 × 60 cm) instead of the 2′,5′-Sepharose column. The column was equilibrated and eluted with buffer C (50 mM Tris–HCl pH 7.5, 300 mM NaCl, 1 mM DTT, 0.1 mM PMSF, 1 mM benzamidine, 1 protease inhibitor tablet/litre) using a flow rate of 1 ml/min. The DPD containing fractions were pooled and concentrated in an Amicon pressure cell to final concentrations of 2–6 mg/ml. At the same time the buffer was exchanged to storage buffer (25 mM HEPES pH 7.5, 10 mM DTT, 10% (v/v) glycerol). Concentrated protein was aliquoted and stored at −20 °C. All purification steps were carried out at 4 °C.

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adjusted as follows: S670A, uracil 0–150 μM, NADPH 0–90 μM; H673N/ Q, uracil 0–25 μM. Concentrations of substrate stock solutions were determined spectroscopically. For comparison one unit of enzyme activity is defined as the amount of enzyme that converts 1 μmol of substrate per hour [13]. For pH studies a composite buffer system containing succinic acid, sodium phosphate and glycine was used at a concentration of 100 mM [18]. Measurements were performed in the pH range 5.0–10.0 in steps of 1 pH unit. All measurements were done at least in duplicate. 2.7. Data processing Initial velocities were averaged and plotted against substrate concentrations. Primary data were fitted using Eq. (1) for Michaelis– Menten kinetics, (2) with substrate inhibition or (3) for ping-pong mechanism analysis. V=

V max ½A K m + ½A 

ð1Þ

V=

V max ½A K m + ½Að1 + ½A = Ki Þ

ð2Þ

V=

V max ½A½B K a ½B + K b ½A + ½A½B

ð3Þ

Secondary parameters Vmax and Km/Vmax were plotted against the pH and the pH-dependent kinetic parameters fitted according to Eqs. (4)–(7). V=

V max H = K1 + K2 = H + 1

V = Km =

V max = K m H = K1 + K2 = H + 1

ð4Þ

ð5Þ

V=

VL 1 + H = K1

ð6Þ

V=

VH 1 + K2 = H

ð7Þ

2.5. Protein concentration determination and UV-Spectroscopy Protein concentrations were determined spectrophotometrically and by the method of Bradford [16]. Absorption spectra from 250– 800 nm were recorded using a JASCO V-650 spectrophotometer. Spectra were measured of 300 μl protein solutions with concentrations of approximately 0.15 mg/ml. DPD concentrations were determined based on the molar extinction coefficient ε of 74 mM−1 cm−1 at 450 nm [17]. Unless otherwise stated DPD concentrations used were determined based on the absorbance at 450 nm.

In Eqs. (1)–(7), V is the initial velocity and Vmax is the maximum velocity. In Eqs. (1)–(3), [A] and [B] are concentrations of substrates and Km, Ka and Kb are Michaelis constants for the respective substrates and Ki is the inhibition constant. In Eqs. (4)–(7), K1 and K2 are acid dissociation constants for enzyme groups; VL and VH are pHindependent values of velocity at low and high pH, respectively; H is defined as 10−pH. Reported standard errors of parameters are derived from residuals of experimental data points to the fits. 2.8. Flavin analysis and iron content determination

2.6. Enzyme assays Enzyme activity was determined at 30 °C by monitoring the decrease in absorbance at 340 nm accompanying the conversion from NADPH to NADP+ as reported previously [13]. A typical reaction mixture contained 100 mM potassium phosphate, 1 mM DTT, 0.5 μg enzyme, and substrates NADPH and uracil in a total volume of 150 μl. The reaction was initiated by the addition of uracil and data were recorded for 5 min in a cell with 1 cm path length. A blank reaction with uracil omitted from the reaction mixture was recorded and subtracted from the reaction measurement. Typical fixed concentrations for one substrate were 10 μM for uracil or 24 μM for NADPH, while the concentration of the other substrate was varied as follows: uracil 1, 2.5, 5, 10 μM; NADPH 3, 4, 6, 12, 24 μM. For some mutants this range was

Identification and quantitation of FAD and FMN in DPD samples were carried out by reverse phase chromatography on an HPLC system similar to the one described previously [8]. Purified enzyme was incubated at 368 K for 20 min in the dark to liberate the flavins. Precipitate was removed by filtration using a 0.22 μm spin filter. Samples were analysed by HPLC separation using a Phenomenex Jupiter 5u C18 300A (250 × 4.6 mm) column with 31%(v/v) methanol in 5 mM ammonium acetate buffer pH 6.0 at a flow rate of 1 ml/min. Flavins were detected at wavelengths 267 and 373 nm (and 230 and 445 nm). Elution profiles were compared with a standard solution containing FAD and FMN in a 1:1 molar ratio. The iron content was determined colorimetrically for WT and mutants C126A and Q156E. Colorimetric analysis was performed

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using the Sigma Diagnostics Iron kit (Sigma Diagnostics) according to the manufacturer's instructions, except that protein was denatured in 1 M perchloric acid. 2.9. EPR spectroscopy EPR spectra (9.5 Ghz) were recorded on a Bruker ESP 300 X-band spectrometer, with an Oxford Instruments ESR9 helium cryostat at 20 K, 3 mW microwave power and 10 mT modulation amplitude. Spin quantitation was performed by double integration of EPR spectra recorded at non-saturating microwave power levels using standard Bruker software, and comparing with a standard solution of 1 mM CuSO4 in 10 mM EDTA. 180 μl of DPD samples at concentrations of approximately 5 mg/ml in standard storage buffer was reduced in the presence of 2 mM dithionite and 2 mM sodium borate, and frozen immediately.

Table 1 Relative activities of WT and mutant DPD. Mutation

Relative activity %

WT C126A Q156E R235A R235K S670A H673N H673Q

100 106 ± 6 0 0 0 73 ± 3 28 ± 2 48 ± 3

Proposed function of mutated residue

Potential [4Fe–4S]-cluster binding [4Fe–4S]-cluster binding FAD binding, proton shuttling FAD binding, proton shuttling Active site loop, uracil binding Active site loop Active site loop

contrast, the mutation of C126 (C126A) did not affect catalytic activity or other kinetic parameters. Determined relative activities are listed in Table 1.

2.10. Figures 3.3. Kinetic properties of mutant DPD Schemes were produced with ISIS/Draw (MDL) and figures with PyMOL [19]. 3. Results 3.1. Expression and purification of wild-type and mutant DPD To simplify the original purification protocol [11], an N-terminal tag comprising six histidines and 11 linker residues was added to porcine DPD by recloning into the vector pET28. All site-specific mutants were derived from this clone and like the wild-type porcine DPD heterologously expressed in E. coli. Expression levels were usually low for both tagged and non-tagged DPD even when high density cultures were obtained using rich auto-induction medium [15]. Co-expression of E. coli iron–sulfur cluster assembly proteins (isc operon) to aid the introduction of [4Fe–4S] clusters did not increase the DPD expression levels (data not shown). Purification of the Histagged enzyme was accomplished in only two steps using a metal affinity (Co2+, Talon) and a 2′,5′-ADP-Sepharose column. However, to obtain milligram quantities of pure DPD large amounts of cells and hence large volumes of lysate are required, the production and processing of which is comparatively time-consuming. Typically 120 g of wet cells yielded between 10 and 20 mg pure WT DPD, while the yield for mutant DPD was about 2–10 times less. The mutants R235A and R235K did not bind to the 2′,5′-ADP-Sepharose column. R235A was further purified using a gel filtration column, yielding enzyme of reasonable but slightly lower purity than enzyme obtained with two affinity chromatography steps. The yield of R235K after the metal affinity chromatography was estimated to be below 0.1 mg and hence no further purification was attempted. Such low amounts were observed repeatedly in different preparations of R235K. All other mutants were purified as the WT enzyme and resulted in purity comparable to that of WT DPD as judged by SDS-PAGE (data not shown). 3.2. Catalytic activity of mutant DPD Initially, all obtained DPD mutants were tested for activity with fixed concentrations of one substrate (10 μM uracil or 24 μM NADPH) while varying the other, to assure that the chosen concentration ranges were either below or only modestly above the previously determined substrate inhibition constants for pig liver DPD of 4 μM and 60 μM for uracil and NADPH, respectively [13]. The active site loop mutants S670A, H673N and H673Q showed catalytic activity comparable to the WT enzyme. Q156E, R235A and R235K showed no activity, even after substantial increase of the substrate concentrations (up to 10 mM uracil and 24 mM NADPH). Enzymatic activity could not be restored by addition and incubation of these mutant enzymes with FAD and FMN. In

Initial velocities were determined with one substrate concentration fixed while varying the other. A plot of the velocities against the varied substrate concentration shows Michaelis–Menten behaviour. The kinetic parameters for WT and mutant DPD were determined using Eqs. (1)–(3). The obtained values for Km and Vmax were independent of the analysis method, i.e. no significant differences were observed. This shows that under the chosen experimental conditions no substrate inhibition occurs, especially with respect to uracil. In Table 2 the kinetic parameters for WT and mutant DPD are shown based on the analysis according to the ping-pong mechanism (Eq. (3)), which should give more reliable fits as compared to Michaelis–Menten analysis. The specific activity for WT DPD is with 8 units/mg about 2 times lower than previously reported [11]. This may be due to different enzyme preparation and protein concentration determination, but it also cannot be excluded that the introduction of a His-tag influences the specific activity. However, this would not compromise a comparative assessment of the kinetic parameters of all tagged enzymes variants. The Km values for tagged WT DPD are 1.2 μM for uracil and 5.7 μM for NADPH comparable to the ones obtained for recombinant non-tagged (Kuracil = 1.0 μM and KNADPH = 6.0 μM) [11] and native pig liver DPD (Kuracil = 1.0 μM and KNADPH = 6.6 μM) [13]. The kinetic parameters for C126A are within the experimental deviations the same as for WT enzyme. Mutation of the active site loop residue H673 to asparagine or glutamine had no influence on the Km of NADPH, while it is about 3 times increased when the nearby located S670 is mutated to alanine. Furthermore, the rate of consumption of NADPH in absence of uracil was about 50% larger in S670A as compared to the WT or other DPD mutants reported here (Km = 46± 15 μM; Vmax = 1.09 ± 0.17 μM/s, i.e. approximately 20% of observed Vmax). A similar behaviour was previously observed for the C671A mutant [17]. The Km for uracil is increased in all three active site loop mutants as compared to the WT. With 37 μM it is highest for the S670A mutant (~30-fold increase). The kcat decreases from S670A to H673Q and H673N. Also the catalytic efficiency as indicated by kcat/Km is decreased in these mutants, with Table 2 Kinetic parameters for WT and mutant DPD. Mutation

WT C126A S670A H673N H673Q

kcat

Km

kcat/Km × 1000

NADPH

Uracil

NAPDH

Uracil

s−1

μM

μM

s−1 μM−1

s−1 μM−1

0.242 ± 0.009 0.256 ± 0.011 0.177 ± 0.004 0.068 ± 0.004 0.117 ± 0.007

5.7 ± 0.5 6.1 ± 0.5 18.8 ± 1.0 7.5 ± 0.8 6.2 ± 0.8

1.2 ± 0.1 1.1 ± 0.1 36.6 ± 1.5 7.0 ± 0.7 4.2 ± 0.6

42 ± 4 42 ± 4 9±1 9±1 19 ± 3

208 ± 22 238 ± 30 5±1 10 ± 1 28 ± 4

B. Lohkamp et al. / Biochimica et Biophysica Acta 1804 (2010) 2198–2206

λ /nm

H673Q showing the least and S670A the most substantial change, with respect to both substrates. Interestingly, in these three mutants the kcat/Km values are similar for both substrates whereas in the WT DPD the efficacy for uracil is about 5 times higher than for NADPH.

0.100

3.5. Spectral properties Besides the peak at 276 nm characteristic for aromatic amino acids the absorption spectrum recorded for WT DPD shows features in the range of 300–500 nm (Fig. 3). Small peaks are observed at 379 and 437 nm and two shoulders at 325 and 450 nm. These features can be attributed to the flavin cofactors and the [4Fe–4S]-clusters and correspond to the ones observed previously for native and recombinant pig liver DPD [8,17]. The spectra for all active site loop mutants (S670A, H673N and H673Q) and of C126A are almost indistinguishable in this Table 3 Summary of pK values from pH dependence of kinetic parameters. V

WT C126A S670A H673N H673Q

V/KNADPH pK2

5.7 ± 0.1 8.7 ± 0.1 Not determined 6.7 ± 0.2 9.3 ± 0.5 6.4 ± 0.1 8.4 ± 0.1 5.4 ± 0.2 9.0 ± 0.2

V/Kuracil

pK1

pK2

pK1

pK2

5.8 ± 0.5

8.8 ± 0.5

6.1 ± 0.2

8.8 ± 0.5

7.0 ± 0.3 5.8 ± 0.4 7.0 ± 0.3

8.6 ± 0.4 8.3 ± 0.5 7.9 ± 0.3

n/a n/a n/a

n/a 8.0 ± 0.2 8.5 ± 0.1

AU

0.075

0.05

0.00 300

WT C126A Q156E R235A S670A H673N H673Q

400

AU

0.10

Kinetic parameters for the DPD reaction were measured over the pH range 5.0–10.0. However, due to the instability of NADPH at acidic and basic pH, data points at pH 10.0 had to be completely omitted from the analysis, while for pH 5.0 only reliably measured data points were taken into account (i.e. duplicate measurements did not differ by more than 50%). Although a different buffer system was used for the pH-dependent determination of kinetic parameters the values for Km and Vmax extrapolated to pH 7.3 correspond well with those determined at fixed pH in phosphate buffer. This and the similarity of pK values presented here and the previously determined ones for native DPD [20] implies that the used buffer system has no or a very limited influence on the kinetic parameters (Table 3). For WT DPD a decrease in the values of V/KNADPH, V/Kuracil as well as Vmax can be observed at both higher and lower pH, indicating that the protonation state of at least two enzyme groups affect the kinetic barriers of the reaction starting with substrate binding and culminating in the first irreversible step in both the free enzyme and the ES complex. The pK values for the two functional enzyme groups determined from the pH dependence of Vmax are pK1 = 5.7 ± 0.1 and pK2 = 8.7 ±0.1, which is in good agreement with the values of 5.6 and 8.8 published earlier for the native pig liver enzyme [20]. The higher acid dissociation constant pK2 is basically unaffected in all three active site loop mutants, but an increase of pK1 to 6.7 and 6.4, respectively, is observed for S670A and H673N. The pH dependence of V/KNADPH gave pK values for WT DPD of 5.8 and 8.8, again comparable to the ones observed for the native pig liver enzyme (5.8 and 8.2). The pK1 for mutants S670A and H673Q was with 7.0 significantly higher than for WT, while the pK2 values remain largely unaffected for all investigated mutants. Also the pK values calculated from the pH dependence of V/Kuracil were in good agreement between WT DPD (6.1 and 8.8) and native pig liver DPD (5.6 and 9.1). Similar pK2 values are obtained for H673N and H673Q, but interestingly, V/Kuracil does not decrease at low pH for these mutants. Therefore the data were analysed with Eq. (7). For the S670A mutant no pK values for V/Kuracil could be calculated since there was no decrease in the pH dependency neither at low nor at high pH within the error of the measurements.

pK1

320 340 360 380 400 420 440 460 480 500

0.15

3.4. pH dependence of kinetic parameters

Mutation

2203

0.050

0.025 500

600

700

800

λ /nm Fig. 3. UV/Vis spectra of WT and mutant DPD. Visible spectra of WT and mutant C126A, S670A, H673N, and H673Q are virtually indistinguishable. The insert shows the difference between the spectra of WT and mutants Q156E and R235A. The characteristic flavin peaks are less pronounced. All spectra have been arbitrary normalised to the WT absorbance at 345 nm.

region from the WT spectrum (Fig. 3), indicating identical flavin and FeS cluster binding. However, mutation of Q156 and R235, which coordinate nFeS2 and FAD, respectively, alters the spectral properties. For Q156E, the main difference is observed in the region around 440 nm where the peak and shoulder merge and level off faster with higher wavelength. Furthermore, the peak at 379 nm is less pronounced. The spectrum for mutant R235A shows even fewer features and an overall lower absorbance. As for Q156E, there is a steep decrease in absorbance beyond the peak at 379 nm, which itself is hardly recognisable. No distinct peak is observed at 440 nm. Hence, for both mutants the spectra indicate a change in the flavin absorbance and/or absorbance related to the FeS clusters. 3.6. Flavin and iron content analysis Flavins were extracted from DPD and analysed on a reverse phase HPLC column. Since flavins usually show four distinct peaks at around 230, 267, 373 and 445 nm elution profiles for these wavelengths were recorded. Due to background noise mainly from the buffer, only the data at 267 and 373 nm were used for analysis. WT DPD shows two peaks corresponding to FAD and FMN respectively. Integral values indicate a 1:1 ratio of the flavins. Analysis of mutant Q156E shows the same elution profile as WT DPD (data not shown), which indicates that this mutation does not affect the binding of FAD (or FMN). Mutant R235A shows only the peak corresponding to FMN in the elution profile (Fig. 4), revealing that this mutant does not bind FAD under the probed conditions. Analysis of the flavin content of mutant R235K was inconclusive due to insufficient amounts of extracted flavin and an unreliably low signal to noise ratio. Iron determination for WT DPD and mutants C126A and Q156E gave similar results of 8 ± 1 iron/mol enzyme subunit. This is only half of the iron expected for the 4 [Fe4–S4] clusters of DPD. This can most likely be attributed to an incomplete release or detection of the iron by the used method, since mass spectrometry of DPD samples indicated complete saturation of the clusters (data not shown). Nevertheless, the results of the iron determination experiments suggest that all investigated DPD variants have the same iron cluster saturation and that absence of one or several FeS clusters is unlikely. 3.7. EPR studies Recorded EPR spectra indicated that efficient reduction of the iron–sulfur centres could only be achieved by treatment with two

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B. Lohkamp et al. / Biochimica et Biophysica Acta 1804 (2010) 2198–2206 WT 267nm WT 373nm R235A 267nm R235A 373nm

0.0025

FAD 0.0020

0.0020

0.0015

FMN

AU

0.0015 0.0010 0.0010 0.0005 0.0005

0.0000 0

1

2

3

4

5

6

7

8

9

0.0000 10

ml Fig. 4. Flavin analysis. Shown are the elution profiles at 267 and 373 nm of the flavins extracted from WT (black and red) and R235A DPD (green and blue). Arrows indicate elution peaks corresponding to FAD and FMN standards. No peaks corresponding to FAD are observed in the R235A mutant indicating a loss of the flavin.

strong reducing agents together, 2 mM dithionite and 2 mM sodium borate. Reduced WT DPD showed an EPR spectrum with well-resolved axial symmetric signals at g|| (2.04 and 2.02) and overlapped at g (1.93–1.94) (Fig. 5). The spectrum and g-values are comparable to the ones observed previously for this enzyme [11,21]. The complex features of the spectrum show at least two different [4Fe–4S]1+clusters. The analysis of the saturation behaviour of the signals at different temperatures (data not shown) allows assignment of the g-values of 2.04 and 1.93 to the centre with higher oxidation potential. This centre could be partly reduced with dithionite (E0 ~ −430 mV) only. The other cluster, with g-values of 2.02 and 1.94, is probably buried deeper in the protein and could only be reduced when

2.03

2.02 2.04

1.94 1.93

3200

a

3600 B, Gauss

b

c 3200

3400

3600

3800

B, Gauss Fig. 5. EPR spectra of recombinant pig DPD. The main panel shows the spectra for reduced WT and mutant DPD and the insert of untreated WT DPD. (a) WT DPD, (b) C126A, and (c) Q156E in presence of 2 mM dithionite and 2 mM sodium borate. EPR spectra for WT and C126A mutant enzyme are identical. No EPR signal is observed for DPD Q156E.

additional reduction with sodium borate (E0 ~ −1811 mV) was performed. The broadening of the peaks for the second type of cluster may indicate that the signal is actually derived from two, very similar clusters. Spectra recorded under non-reducing and oxidizing conditions showed a minor signal for a high potential iron–sulfur cluster (e.g. [3Fe–4S]1+). The signal was estimated to be maximally 3% of the reduced form. This may be explained by degradation and loss of an iron ion by one of the [4Fe–4S] clusters and considered as an artefact. EPR spectra recorded for C126A are almost identical to those for WT. DPD mutant Q156E only shows the very weak signal characteristic for a radical that is present in all DPD EPR spectra. No FeS cluster signal could be observed under reducing conditions, which suggests that the clusters cannot be reduced in this mutant. EPR spectra recorded under non-reducing and oxidizing conditions look similar to the ones of WT DPD. 4. Discussion 4.1. Iron–sulfur cluster binding It was recently proposed that the N-terminal half of DPD, the SudA subunit of Pyrococcus furiosus sulfide dehydrogenase, and the β-subunit of bacterial and non-photosynthetic eukaryotic glutamate synthases (GltS) are members of a novel family of FAD-containing NADPH oxidoreductases, which transfer electrons to an acceptor protein or domain through [4Fe–4S] clusters of low to very low potential [22]. All contain two cysteine-rich patterns providing the cluster-coordinating residues in an N-terminal extension (domain I of DPD) that is fused to an adrenodoxin reductase-like module binding FAD and NADPH (domains II and III) [22]. For DPD the midpoint potential was experimentally determined to be ~−440 mV [21], and cysteines 91, 126, 130 and 136 would be available for binding nFeS2 [21]. However, despite its strict conservation in DPD sequences C126 does not serve as a cluster ligand [12] (Fig. 2c). Instead, the fourth iron is coordinated by Q156. In SudA and GltS sequences an invariant glutamate replaces the DPD-Q156, while the cysteine corresponding to C126 is often replaced by threonine. It has therefore been proposed that one of the two FeS clusters in these enzymes is a [3Fe–4S] [22]. Alternatively, the invariant glutamate replacing the DPD-Q156 in these homologous proteins could serve as a fourth ligand for coordination of a [Fe4–S4] cluster. Our mutational studies revealed that the conservation of the cysteine at position 126 reflects limitations in available space for the side chain rather than functional importance for DPD. The kinetic properties and the EPR spectrum of the mutant are unaltered compared to WT DPD, confirming that C126 is not involved in nFeS2 binding and has no significant influence on its redox potential. Mutation of Q156 to glutamate resulted in an inactive enzyme. nFeS2 cannot be reduced even by the harshest conditions used in the EPR studies. The redox potential of this cluster is most likely significantly lowered due to the introduction of a negative charge on this cluster ligand, making NADPH via FAD unable to reduce it. Since only half of the cluster metals could be detected in the iron determination experiments, it cannot entirely be excluded that Q156E lost nFeS2 or at least one of its metal ions. However, detection of the same iron content for WT DPD, comparable protein stability as well as the absence of an EPR signal under oxidizing conditions characteristic for [3Fe–4S] clusters argue against it. The absence of any FeS-cluster signal in the EPR spectrum of the Q156E mutant is in agreement with the arrangement of the prosthetic groups in DPD: nFeS2 is the first cluster to be reduced. In the WT enzyme but not Q156E, electrons are transferred from here to the one or two cluster(s) next in line. Since the ligation of clusters nFeS1 and cFeS1 with four cysteine residues is similar, both may contribute to the broadened second peaks in the recorded spectra. Hence, cluster cFeS2 at the pyrimidine binding site

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seems to be the cluster which is not reducible under the experimental conditions of the EPR studies. 4.2. FAD binding Recurrent features of flavoenzymes catalyzing dehydrogenation reactions are the presence of a positively charged group near the N1 locus of the FAD-isoalloxazine ring, and the interaction of the N5 locus with a hydrogen-bond donor [23]. In contrast to the interactions with the N1 locus, the role of the hydrogen bond to the N5 locus has not yet been well studied. Fraaije and Mattevi [23] proposed that it stabilizes the oxidized state of the cofactor as the bond becomes energetically less favourable upon protonation of N5 due to hydride transfer from NADPH. In DPD, the guanidinium group of R235, one of the residues found mutated in patients with DPD deficiency that present with thymine-uraciluria or experienced severe 5-fluorouracil toxicity [24,25], is placed in close proximity to the N5 locus. It hydrogen bonds to FAD-O4, but can also form a hydrogen bond with the N5-atom. Its location and interaction with D346, which in turn is connected to E376 and the bulk solvent, would make R235 prone to be involved in shuttling the N5-proton away after electron transfer to the FeS clusters. However, an arginine is not expected to be a good proton abstractor. The exchange of R235 to alanine resulted in an inactive enzyme lacking FAD. Incubation with FAD did not restore enzyme activity. The loss of the cofactor also compromises enzyme stability, since a higher tendency for aggregation was observed (data not shown). Unexpectedly, presence of a lysine at position 235 does not even partly compensate for the loss of the arginine but has even more deleterious effects. The low expression levels of the R235K mutant suggest that it is very unstable. No residual enzyme activity is detected, most likely because this mutant is also unable to bind FAD. The severity of the effects caused by the point mutations shows that the presence of an arginine at position 235 is first and foremost essential for the binding of FAD, either via direct interactions with the cofactor or by playing a crucial role in the construction of its binding site. Further studies regarding its influence on the electronic state of the cofactor or regarding potential other roles in catalysis will require a R235 DPD mutant which is able to retain FAD. 4.3. Pyrimidine binding and active site closure In “open active site loop” crystal structures of DPD, all loop residues beyond S670 are fully solvent-exposed and partly disordered. S670 is hydrogen-bonded to N668, a substrate binding residue, and also weakly to the substrate itself [12]. Hence, S670 may initially be involved in substrate recognition. In “closed loop” structure, S670 anchors one loop end via hydrogen bonds with the backbone oxygen atom of the FMN-binding K709 and the side chain of the substrate binding N736, respectively (Fig. 2e). Additional loop anchor points are H673 forming a hydrogen bond to E611, M675 engaging in hydrophobic interactions with non-loop residues, and E677 forming hydrogen bonds to residues 935 and 936. The latter two seem less important, because the second half of the loop retains a higher degree of flexibility also in the closed state. The role of H673 as anchor point explains why in DPD crystal structures determined at pH 4.7 open loops were observed even when a substrate analogue was bound [12,14]: due to its protonation loop closure would introduce an uncompensated positive charge into the active site and cause close contacts to P672. To further investigate the importance of S670 and H673 for the stabilization of a closed loop we targeted both residues for site-directed mutagenesis. Previous studies on the pH dependency of kinetic parameters of DPD identified a group with a pK of ~ 6.5, which needs to be unprotonated for optimal catalytic activity but is not essential [20]. Based on the absence of a low pK with respect to the pyrimidine substrate for the H673Q and H673N mutants we propose that H673 accounts for this group, at least in part. The residual activity of the mutant enzymes shows that H673 is not essential but important for

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catalysis to proceed optimally. However, the unchanged low pK with respect to V for H673Q (pertinent for the ES complex) and V/KNADPH for H673N (pertinent for the free enzyme) indicates that there is a second enzyme group, presumably at the NADPH binding site, that needs to be unprotonated for optimal catalysis. The reduced kcat and increased Km for the substrate uracil of both mutants are likely to reflect structural rearrangements in the active site required for substrate binding and loop closure. Due to the similar side chain lengths of glutamine and histidine the hydrogen bond between residue 673 and E611 in the closed loop state could be preserved in H673Q. However, the resulting bond geometry is not favourable. In case of H673N the smaller side chain of N673 would necessitate larger structural rearrangements to achieve loop closure, which may explain the lower catalytic activity and higher Km for uracil compared to H673Q. Nevertheless, full or partial closure of the active site upon substrate binding seems to be achieved as significant residual activity is observed. Although effects on Km cannot directly be translated into changes in substrate affinity, the almost 40-fold increase in the Km for uracil for S670A strongly suggests a major involvement of the residue in processes leading to formation or dissociation of the ES(uracil) complex, i.e. in pyrimidine binding and recognition. Crystal structures of DPD complexes [12,14] do not reveal strong interactions of S670 with substrate analogues. Thus, the elevated Km may instead be explained with a stabilisation and hence higher population of the closed loop conformation in the mutant enzyme, which prevents access to the substrate binding site. The concomitant increase in Km for NADPH is remarkable considering that the NADPH binding site is approximately 60 Å away from the S670A mutation site. Furthermore, the rate of NADPH oxidation in absence of the pyrimidine substrate is increased, as it was previously observed for the C671A mutant [17]. This indicates that there is a mechanism in place that prevents futile NADPH oxidation in absence of pyrimidine in the native enzyme. Electron flow from the NADPH should only occur after substrate binding and closure of the active site. Consequently, closure should either be hampered or not be able to trigger the electron flow when the pyrimidine site is empty. The available data allow us to speculate on a possible scenario how this could be achieved. Since the mutation of H673 does not affect the oxidation rate or the Km for NADPH it seems that specific contacts of S670 (and possibly C671), not the sole event of active site loop closure, are important for the communication between the ligand binding sites. An important function of S670 could be to prevent proper or complete loop closure and hence electron flow when the active site is unoccupied, possibly via the hydrogen bond to N668. This hydrogen bond will be weakened upon substrate binding by interactions of both residues with the pyrimidine, allowing complete active site closure with S670 now forming hydrogen bonds to K709 and N736. The switch in hybridization of atoms C5 and C6 from sp2 to sp3 upon reduction of the substrate, which makes the ring non-planar, may aid loop opening and product release by pushing adjacent residues aside. Loss of the serine hydroxyl group in S670A would prevent stabilization of an open loop conformation. Full closure can occur even without the substrate, explaining the higher NADPH oxidase activity in absence of pyrimidine and hence the uncoupling of the electron transfer from the catalytic cycle. But how is triggering of the electron flow from NADPH achieved on a molecular level? An attractive hypothesis would be that sequestration of the active site from the bulk solvent together with specific substrate and active site loop interactions affects the electronic state of FMN and/or a FeS cluster, possibly the one which could not be reduced in the EPR studies. Adjustments of the redox potential would enable it to serve as an electron sink and thus ensure a rapid electron flux through the enzyme, but only when pyrimidine is bound. Nevertheless this does not provide an explanation for the apparent higher Km for NADPH for the S670A mutant. It seems unlikely that the mutation causes significant structural changes in the NADPH binding site, because of the large distance between both loci and because no differences have been observed between the

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NADPH binding sites in DPD structures with open and closed active site loop obtained at the same crystallization pH [14]. On the other hand, the increase in the lower pK determined from the pH dependence of V/ KNADPH for the mutants S670A and H673Q supports a model for the communication between the active sites that relies on long-range transmitted conformational changes. According to previous studies on native pig liver DPD the titrated functional group is not NADPHassociated but must be an enzyme group that is likely involved in maintaining the structural integrity of the NADPH binding site. [20]. In conclusion, further investigations into the exact mechanism of the coupling of the electron transfer with active site loop movement are required to explain this and other observations. Nevertheless to our knowledge this is the first time an uncoupling of electron transfer from the reduction site in combination with a negative allosteric influence on the co-enzyme binding is observed. Acknowledgements We would like to acknowledge the financial support from Cancerfonden, the Swedish Research Council and Wenner-Grenska Samfundet. References [1] C. Wasternack, Degradation of pyrimidines and pyrimidine analogs—pathways and mutual influences, Pharmacol. Ther. 8 (1980) 629–651. [2] T.W. Traut, M.E. Jones, Uracil metabolism-UMP synthesis from orotic acid or uridine and conversion of uracil to beta-alanine: enzymes and cDNAs, Prog. Nucleic Acid Res. Mol. Biol. 53 (1996) 1–78. [3] W.E. Hull, R.E. Port, R. Herrmann, B. Britsch, W. Kunz, Metabolites of 5-fluorouracil in plasma and urine, as monitored by 19F nuclear magnetic resonance spectroscopy, for patients receiving chemotherapy with or without methotrexate pretreatment, Cancer Res. 48 (1988) 1680–1688. [4] R.B. Diasio, T.L. Beavers, J.T. Carpenter, Familial deficiency of dihydropyrimidine dehydrogenase. Biochemical basis for familial pyrimidinemia and severe 5-fluorouracilinduced toxicity, J. Clin. Invest. 81 (1988) 47–51. [5] A.B. van Kuilenburg, Dihydropyrimidine dehydrogenase and the efficacy and toxicity of 5-fluorouracil, Eur. J. Cancer 40 (2004) 939–950. [6] J.S. de Bono, C.J. Twelves, The oral fluorinated pyrimidines, Invest. New Drugs 19 (2001) 41–59. [7] Z.H. Lu, R. Zhang, R.B. Diasio, Purification and characterization of dihydropyrimidine dehydrogenase from human liver, J. Biol. Chem. 267 (1992) 17102–17109. [8] B. Podschun, G. Wahler, K.D. Schnackerz, Purification and characterization of dihydropyrimidine dehydrogenase from pig liver, Eur. J. Biochem. 185 (1989) 219–224.

[9] T. Shiotani, G. Weber, Purification and properties of dihydrothymine dehydrogenase from rat liver, J. Biol. Chem. 256 (1981) 219–224. [10] D.J. Porter, W.G. Chestnut, L.C. Taylor, B.M. Merrill, T. Spector, Inactivation of dihydropyrimidine dehydrogenase by 5-iodouracil, J. Biol. Chem. 266 (1991) 19988–19994. [11] K. Rosenbaum, B. Schaffrath, W.R. Hagen, K. Jahnke, F.J. Gonzalez, P.F. Cook, K.D. Schnackerz, Purification, characterization, and kinetics of porcine recombinant dihydropyrimidine dehydrogenase, Protein Expr. Purif. 10 (1997) 185–191. [12] D. Dobritzsch, G. Schneider, K.D. Schnackerz, Y. Lindqvist, Crystal structure of dihydropyrimidine dehydrogenase, a major determinant of the pharmacokinetics of the anti-cancer drug 5-fluorouracil, EMBO J. 20 (2001) 650–660. [13] B. Podschun, P.F. Cook, K.D. Schnackerz, Kinetic mechanism of dihydropyrimidine dehydrogenase from pig liver, J. Biol. Chem. 265 (1990) 12966–12972. [14] D. Dobritzsch, S. Ricagno, G. Schneider, K.D. Schnackerz, Y. Lindqvist, Crystal structure of the productive ternary complex of dihydropyrimidine dehydrogenase with NADPH and 5-iodouracil. Implications for mechanism of inhibition and electron transfer, J. Biol. Chem. 277 (2002) 13155–13166. [15] F.W. Studier, Protein production by auto-induction in high density shaking cultures, Protein Expr. Purif. 41 (2005) 207–234. [16] M.M. Bradford, A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal. Biochem. 72 (1976) 248–254. [17] K. Rosenbaum, K. Jahnke, B. Curti, W.R. Hagen, K.D. Schnackerz, M.A. Vanoni, Porcine recombinant dihydropyrimidine dehydrogenase: comparison of the spectroscopic and catalytic properties of the wild-type and C671A mutant enzymes, Biochemistry 37 (1998) 17598–17609. [18] J. Newman, Novel buffer systems for macromolecular crystallization, Acta Crystallogr. D 60 (2004) 610–612. [19] W.L. DeLano, The PyMOL Molecular Graphics System, in, DeLano Scientific, Palo Alto, CA, USA, 2002. [20] B. Podschun, K. Jahnke, K.D. Schnackerz, P.F. Cook, Acid base catalytic mechanism of the dihydropyrimidine dehydrogenase from pH studies, J. Biol. Chem. 268 (1993) 3407–3413. [21] W.R. Hagen, M.A. Vanoni, K. Rosenbaum, K.D. Schnackerz, On the iron–sulfur clusters in the complex redox enzyme dihydropyrimidine dehydrogenase, Eur. J. Biochem. 267 (2000) 3640–3646. [22] M.A. Vanoni, B. Curti, Structure-function studies on the iron–sulfur flavoenzyme glutamate synthase: an unexpectedly complex self-regulated enzyme, Arch. Biochem. Biophys. 433 (2005) 193–211. [23] M.W. Fraaije, A. Mattevi, Flavoenzymes: diverse catalysts with recurrent features, Trends Biochem. Sci. 25 (2000) 126–132. [24] A.B.P. van Kuilenburg, J. Meijer, D. Dobritzsch, B. Lohkamp, W. Ruitenbeek, J. Roelofsen, N.G.G.M. Abeling, M. Duran, C. Buzing, Identification of two novel mutations C79X and R235Q in the dihydropyrimidine dehydrogenase gene in a patient presenting with hematuria, Nucleosides Nucleotides Nucleic Acids 27 (2008) 809–815. [25] P. Vreken, A.B. Van Kuilenburg, R. Meinsma, A.H. van Gennip, Dihydropyrimidine dehydrogenase (DPD) deficiency: identification and expression of missense mutations C29R, R886H and R235W, Hum. Genet. 101 (1997) 333–338. [26] K.D. Schnackerz, D. Dobritzsch, Y. Lindqvist, P.F. Cook, Dihydropyrimidine dehydrogenase: a flavoprotein with four iron–sulfur clusters, Biochim. Biophys. Acta 1701 (2004) 61–74.