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1 Department of Biological Chemistry, Institute of Molecular Biology, University of Copenhagen, DK-1307, Copenhagen K, Denmark
2 Centre for Crystallographic Studies, Department of Chemistry, University of Copenhagen, DK-2100, Copenhagen 0, Denmark
3 Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109-0606 USA
Reprint requests to: Kaj Frank Jensen, Department of Biological Chemistry, Institute of Molecular Biology, University of Copenhagen, Sølvgade 83H, DK 1307 Copenhagen K, Denmark; e-mail: kfj{at}mermaid.molbio.ku.dk; fax: +45 3532 2040.
(RECEIVED June 20, 2002; FINAL REVISION August 14, 2002; ACCEPTED August 15, 2002)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.0220302.
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Keywords: Nucleotide synthesis; oligomerization; quaternary structure; dissociation kinetics; oxido-reductase; flavin
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Introduction |
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Two types of Class 1 DHODs, 1A and 1B, have been defined. The Class 1A DHODs are found in Lactococcus lactis (Nielsen et al. 1996b), Enterococcus faecalis (Marcinkeviciene et al. 2000), anaerobic yeast (Nagy et al. 1992; Jordan et al. 2000), and some protozoan parasites (Pascal et al. 1983; Gao et al. 1999). They are dimeric proteins that use fumarate, quinones, and O2 as electron acceptors (Rowland et al. 1997a; Björnberg et al. 1999, 2001). The DHODs of Class 1B are prevalent in Gram-positive bacteria such as L. lactis (Nielsen et al. 1996a), E. faecalis (Marcinkeviciene et al. 1999), Bacillus subtilis (Kahler et al. 1999), and Clostridium oroticum (Argyrou et al. 2000). They are heterotetrameric proteins (PyrDb2 PyrK2) which utilize a subunit belonging to the ferredoxin reductase family (PyrK) to channel the electrons from the oxidation of DHO to NAD+ (Nielsen et al. 1996a; Rowland et al. 2000).
We have investigated dihydroorotate dehydrogenase A (DHODA) from L. lactis as a representative for the Class 1A DHODs. This enzyme contains one molecule of FMN bound to each subunit (Nielsen et al. 1996b), and the crystal structure of the protein is known (Rowland et al. 1997b; 1998). Each subunit folds into an (
/ß)8-barrel structure, and the active site is formed by several loops that protrude from the top of the barrel. When bound in the active site, the substrate dihydroorotate is stacked with the isoalloxazine ring of FMN in an orientation favorable for hydride-transfer from the C6 position of DHO to the N5-position of FMN (Rowland et al. 1998; Fraaije and Mattevi 2000; Palfey et al. 2001). The active site base, Cys130, initiates the reaction by abstracting the 5-proS proton from DHO (Björnberg et al. 1997, 2001), to facilitate hydride transfer to the flavin. The pKa of the 5-proS proton of DHO is 20–21 (Argyrou and Washabaugh 1999). However, many tight hydrogen bonds between the substrate and totally conserved asparagine side chains of the enzyme, notably Asn132 and Asn193, which interact with the O4 position of DHO, may serve to increase the acidity of the 5-proS proton in the enzyme-substrate complex (Fraaije and Mattevi 2000; Björnberg et al. 2001). Furthermore, the reaction may be facilitated by the interaction of Lys43 with the flavin, which stabilizes the negative charge that develops on the isoalloxazine ring upon hydride transfer (Björnberg et al. 1997; Jiang et al. 2000).
The two subunits of DHODA are related by a noncrystallographic twofold axis. Twenty-eight amino acid residues from each subunit form the dimer interface through hydrophobic and hydrophilic interactions. There are two intersubunit salt bridges formed between E206 of one subunit and K296 of the other subunit (Fig. 1
). E206 is part of a small 310-helix, and K296 is placed at the end of helix 8. These two residues are completely conserved within the Class 1A DHODs. A rather hydrophobic cavity (109 Å3) is formed in dimer interface around the center of the twofold axis and contains two water molecules (Rowland et al. 1997b).
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). To test the importance of these salt bridges for activity, we changed the residues involved in forming the salt bridges to either alanine or the amino acid of opposite charge, that is, lysine to glutamate or glutamate to lysine. The following amino acid replacements were constructed: E206A, E206K, K296A, K296E, and also the double replacement mutant E206K-K296E, in which the polarity of the salt bridge was inverted. The genes encoding these mutations contained on the inducible multicopy vector pUHE23-2 were introduced into the E. coli strain SØ6645, which lacks the chromosomal pyrD gene and requires an exogenous source of pyrimidine due to lack of dihydroorotate dehydrogenase (Nielsen et al. 1996b).
Complementation and analysis in crude extracts
Bacterial strains carrying plasmids with mutated as well as wild-type versions of pyrDa were spread as single colonies on agar plates in the absence of uracil (without induction). When plates were incubated at 37°C, only the wild-type pyrDa gene gave rise to colony formation; none of the mutant plasmids showed any complementation. The response was more differentiated at 25°C, as the single mutant E206A relieved the pyrimidine requirement of SØ6645 as well as the wild-type gene on pFN1, and the single mutant K296A and the double mutant E206K-K296E gave rise to fair growth in the absence of uracil. The two single mutants, E206K and K296E, complemented the pyrD deletion of the host extremely poorly if at all (Table 1).
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5 min before an assay at 25°C, E206A and K296A retained all and 10% of the activity, respectively, whereas the other mutants lost all of their activity. These results show that the salt bridge between Glu206 and Lys296 is not required for the enzymatic activity of DHODA, but is important for the stability of the protein at increased temperature. The conversion of one of the amino acids of the salt bridge to opposite charge, as in the E206K and K296E single mutants, is very harmful to the function of DHODA, probably due to electrostatic repulsion over the subunit interface, and this effect is partially relieved by reestablishing a possibility of forming a salt bridge in opposite orientation to the E206K-K296E double mutant enzyme.
E206K-K296E mutant and wild-type DHODA lose activity upon dilution
The mutant E206K-K296E was purified as described in Materials and Methods. The activity was characterized by the use of DCIP (50 µM) as electron acceptor and by recording the disappearance of absorption at 600 nm as a measure of the initial reaction velocity. The kinetic parameters (KMapp = 7.2 ± 0.5 µM for DHO and a kcatapp = 22.4 ± 0.3 s-1) were very similar to those of wild-type DHODA (Björnberg et al. 1997). In addition, the affinity of the mutant enzyme for the product orotate (KD = 37 ± 2 µM) was similar to that of the wild-type enzyme (KD = 34 ± 2 µM) as revealed by recording the red-shift of the absorption spectrum during titration of enzyme solutions with orotate (Björnberg et al. 1997).
However, when the enzymes were diluted in 0.1 M Tris-HCl pH 8.0 prior to assay, the specific activity of the mutant enzyme gradually decreased as a function of time and approached a constant value (Fig. 2B). The loss of activity could be fitted to first-order exponential decay function
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, the rate of inactivation did not depend much on the protein concentration. The inactivation rate constant kobs was close to 0.1 min-1, corresponding to a half-life of approximately 6 min. However, the level of remaining activity (Ar) was strongly dependent on the protein concentration, as shown by the individual inactivation curves in Figure 2B. Such a behavior is most easily explained by assuming that the active dimeric mutant enzyme undergoes a slow and reversible dissociation into inactive monomers according to the following scheme, which also is supported by gel-filtration analyses (see below):
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A number of kinetic parameters relating to the dissociation reaction could be determined from the decay curves in Figure 2, and these are given in Table 2.
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. The resulting saturation curve could be used to calculate the dissociation constant (KD) for the dimer to monomer dissociation reaction, which is given in the legend to the figure. From these data it appears that the dissociation constant KD = (k+1/k-1) for the E206K-K296E mutant enzyme in 0.1 M Tris-HCl pH 8.0 is about 37 µM at 25°C, and that the rate constant for the dissociation reaction k+1 was 0.093 min-1, assuming that kobs equals k+1 for the most dilute enzyme samples, which undergo almost complete dissociation. In turn, these parameters allow for calculation of a second-order rate constant k-1 for association of inactive monomers to active dimers of 0.0025 µM-1 min-1.
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). The first-order rate constant for the inactivation k+1 was 0.03 min-1, corresponding to a half life t
of about 22 min and independent of protein concentration to a first approximation. A measurement of the remaining enzyme activities at different enzyme concentrations (Fig. 3) revealed the constant for dissociation of active dimers to inactive monomers, KD = 0.26 µM. The dissociation rate constant k+1 was 0.024 min-1 (assumed again to equal kobs at the most dilute concentration of enzyme), and thus the second-order rate constant k-1 for association of inactive monomers to active dimers will be 0.043 µM-1 min-1. Thus, the rate of both the dissociation of active dimers to inactive monomers and the reverse association reaction was affected by the E206K-K296E mutations in the enzyme, but the association rate was more affected than the dissociation rate (Table 2).
Effect of salt and the product orotate on the inactivation of E206K-K296E mutant DHODA
The inactivation of the E206K-K296E mutant DHODA was influenced by the salt concentration of the dilution buffer (Fig. 2C). The presence of 0.15 M NaCl reduced the dissociation rate constant k+1 (kobs) to 0.026 min-1 and the equilibrium constant for the dissociation process KD to 3.5 µM (Fig. 3). The association rate constant k-1 was increased to 0.0074 µM-1 min-1, and thus the salt influenced both the rates of association of monomers and dissociation of dimers (Table 2). As shown in Table 3, the product of the enzymatic reaction, orotate, also protected the enzyme against inactivation upon dilution, although the kinetic behavior was not analyzed in detail.
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Gel-filtration behavior of mutant and wild-type DHODA
The wild-type and E206K-K296E mutant enzymes behaved very similarly during gel-filtration chromatography on a Superose 12 column and eluted as single symmetric peaks of a size similar to that of bovine serum albumin (Mr 67.000, i.e., corresponding to the size of a dimeric DHODA molecule) when they were loaded on the column at high concentration, 3.9 mg/mL (114 µM) in the 200 µL sample (data not shown). When the concentration of enzyme in the samples was reduced to 0.14 mg/mL (4 µM), the wild-type enzyme retained its elution behavior, but the mutant enzyme developed a pronounced shoulder on its elution profile corresponding to the occurrence of lower-molecular-weight species in the sample (Fig. 4A). This shoulder was not seen if the gel filtration was made in Buffer A containing 1 M NaCl (data not shown), and it became less pronounced when 600 µM orotate was included in the enzyme samples that were loaded on the column (Fig. 4B). This behavior during gel filtration is in agreement with the notion that the dimeric E206K-K296E double mutant enzyme has an increased tendency to dissociate to (inactive) monomers upon dilution, and that factors (NaCl and orotate) which protect the enzyme from loss of activity during dilution also stabilize the dimeric form of the enzyme.
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), by the increasing (and saturating) specific activity as a function of increasing protein concentration (Fig. 3), and by the behavior of mutant and wild-type enzyme during gel filtration (Fig. 4), as well as the recovery of enzymatic activity upon addition of salt or orotate. The dissociation reaction was more pronounced in the double mutant E206K-K296E in which the polarity of two salt bridges between the two subunits of the dimer (Rowland et al. 1997b) have been inverted. The equilibrium constants for the dimer to monomer dissociation reaction could be estimated both from the end points of the kinetic analyses shown in Figure 2, that is, from the initial activity Ao and the equilibrium activity Ar, and from analysis of equilibrium specific activity as a function of protein concentration (Fig. 3). The values are given in Table 2. The crude determinations of KD from Ao and Ar by the first method tended to generate smaller values than were found by the second method. We ascribe this (minor) discrepancy to the fact that the first method depends on the assumption that all enzyme exists as dimers prior to dilution. This may not be the case, and hence Ao is likely to be underestimated—an error that leads to underestimation of the concentration of monomers at equilibrium and consequently also to (falsely) low values of KD. Another qualification to our calculations is our assumption that the monomers are totally devoid of activity. As illustrated in Figure 2B, this may not be the case, since the decay curve for the most dilute solution (0.1 µM) of the E206K-K296E double mutant enzyme approached an equilibrium specific activity of 1.6 U/mg, when close to zero specific activity was expected from the KD value (37 µM). Therefore, we cannot exclude the possibility that the monomer has an enzymatic activity, which corresponds to 5%, or less, of that of the dimeric enzyme.
Despite the qualifications described above, it is clear that the double E206K-K296E mutation has destabilized the dimeric form of DHODA by a factor of approximately 100, and that the rate of association of monomers to dimers was more strongly affected (40-fold reduced) than the dissociation of dimers to monomers, which occurred 3–4 times faster for the mutant enzyme than the wild type. The fact that the dimeric form was stabilized, rather than destabilized, by sodium chloride indicates that the hydrophobic interactions between dimers are more important than the electrostatic forces, which are expected to be weakened by the presence of salt.
The dissociation constant KD for the dimer to monomer conversion reaction for the wild-type enzyme was estimated to be 0.26 µM, corresponding to a 
G°d = 8.98 kcal mol-1 for the wild-type DHODA and 37 µM for the E206K-K296E mutant enzyme, corresponding to a G°d = 6.04 kcal mol-1. This means that inversion of the two salt bridges in the E206K-K296E mutant enzyme has destabilized the dimer formation to 
G°d = 2.94 kcal mol-1. The architecture of the intersubunit salt bridge in the wild-type enzyme is shown in Figure 1B. (Unfortunately, we have not been able to elucidate the structure of the mutant enzyme, because we were unable to obtain crystals despite several attempts.) It is clear that Lys296 and Glu206 are part of a complex network of hydrogen bonds that includes the side chain -OH of Tyr298, the backbone carbonyl oxygen of Asp204 and several water molecules. Most of these hydrogen bonds are likely to be perturbed in the double mutant enzyme, and it is therefore not possible to assign the decrease in stability of the dimeric form of the mutant enzyme to a single intermolecular interaction.
It can be concluded, however, that the salt bridge is not needed for the catalytic function of DHODA, because the mutant enzymes, in which one of the residues involved in salt bridge formation was replaced by alanine, were also catalytically active, although they tended to dissociate and precipitate under certain conditions. In addition, the loss of enzymatic activity upon dissociation was not due to loss of the prosthetic FMN from the monomers, and thus our results show that the catalytic function of the two active sites is dependent on the presence of the neighboring subunit of the dimer, which perhaps protects the active sites against access of solvent. Although the two active sites are located at some distance from the subunit interface, the loops that cover the active sites are in contact with the neighboring subunit in both the dimeric and the heterotetrameric Class 1 dihydroorotate dehydrogenases DHODA (Rowland et al. 1997b; 1998) and DHODB (Rowland et al. 2000). Removal of the neighboring subunit is likely to influence the conformation of these loops so that they no longer adopt a structure suited for optimal enzyme function. In the monomeric Class 2 DHODs from human (Liu et al. 2000) and Escherichia coli (Nørager et al. 2002), an N-terminal domain responsible for communication with the respiratory quinones and attachment to membranes interacts with the DHOD core domain at a region similar to the one which is involved in dimerization in the Class 1 enzymes.
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General instrumentation
A Zeiss Specord S10 diode-array photometer was used for the enzyme assays and absorption spectra.
Construction of mutant DHODAs
The mutations were introduced in the pyrDa gene using plasmid pFN1 (Nielsen et al. 1996b) as template for polymerase chain reactions (PCRs) with mutant primers as described by Björnberg et al. (1997). The mutated pyrDa genes were cloned in pUHE23-2 (Deuschle et al. 1986) to produce plasmids similar to expression vector for the wild-type enzyme, pFN1 (Nielsen et al. 1996b), and DNA sequencing revealed that they contained no other mutation than the desired one(s). The plasmids were transformed into the Escherichia coli strain SØ6645, which lacks its native dihydroorotate dehydrogenase due to a deletion of the chromosomal pyrD gene and requires exogenous pyrimidine for growth. The strain SØ6645 also contains an F‘lacIq episome, which causes overproduction of the lacI repressor and keeps transcription of the cloned pyrDa gene at a low level until induction by addition of isopropyl-ß-D-thiogalactoside (IPTG).
Complementation of a pyrD mutation
The ability of the mutant pyrD gene harbored in plasmid pUHE23-2 to complement the chromosomal pyrD deletion was tested by plating the strain SØ6645 transformed with the plasmids on agar plates containing glucose (0.2%), casamino acid (0.2%), and thiamine (2 µg/mL) and incubating them at either 25°C or 37°C. The size of colonies was inspected after approximately 24 and 48 h and compared with similar control plates also containing uracil (20 µg/mL). The complementation test was carried out in the absence and in the presence of the inducer IPTG (50 µM), but only the results in the absence of induction were used, because the induction of DHODA mutant—as well as wild-type enzymes—strongly inhibited colony growth even in the presence of uracil.
Crude extract analyses
Ten-mL cultures of the transformed strains were grown in LB-broth medium (Miller 1972) at 25°C or 37°C, with vigorous shaking. The cultures were induced at an OD436 of about 0.5 by addition of IPTG (0.75 mM) and grown to stationary phase overnight. The cultures were harvested by centrifugation for 10 min at 5000 rpm at 5°C (Sorvall SS-34). Cells were suspended in 1 mL of Buffer A (50 mM potassium phosphate, pH 6.0, 10% glycerol, and 1 mM EDTA) and disrupted by ultrasonic treatment during cooling to produce a crude extract. The extract was further fractionated by centrifugation as above. The pellets and supernatant as well as the crude extracts were analyzed for activity using the DCIP assay described below and for protein content by 12.5% SDS-PAGE. Protein concentrations were determined by the Lowry method.
Purification of wild-type and E206K-K296E double mutant DHODAs
Purification of wild-type DHODA was carried out as described (Nielsen et al. 1996b). The E206K-K296E double mutant enzyme was purified similarly, but the transformed strain was grown at 25°C. During purification, chromatography on the hydroxylapatite column was omitted and, instead, the enzyme solution was subjected to chromatography twice on a DE52-column. It appeared to be 
95% pure by SDS gel electrophoresis. The concentration of purified enzymes was estimated from the absorption spectra using the extinction coefficient of FMN of 11.5 mM-1 cm-1 at 456 nm as described (Björnberg et al. 1997).
Enzymatic activity
The oxidation of DHO was measured with DCIP as electron acceptor and monitored by recording the decrease of absorption at 600 nm due to the reduction of dichloroindophenol (DCIP), 
600 = 20 mM-1 cm-1 (Karibian 1978; Björnberg et al. 2001). The UV lamp was turned off during these measurements to avoid photobleaching of DCIP. The assay mixture (1 mL) contained 0.1 M Tris-HCl, pH 8.0, 50 µM DCIP, 0.5 mM (S)-DHO and was started by adding the enzyme. One unit corresponds to 1 µmol of orotate produced by the enzyme per min at 25°C.
Gel filtration
Gel-filtration FPLC using a BioRad BioLogic Workstation was performed on a 25-mL Superose-12 column (Pharmacia) at room temperature. A volume of 200 µL enzyme solution was loaded on the column at a flow-rate of 0.5 mL/min, and fractions of 0.5 mL were collected. Buffer A (50 mM potassium phosphate, pH 6.0, 10% glycerol, and 1 mM EDTA) was used as running buffer in most cases, but in some cases 1 M NaCl was included in the buffer, or 0.6 mM orotate was included in the applied enzyme sample.
Loss of enzymatic activity caused by dilution
A concentrated enzyme stock (5–10 mg/mL, 150–300 µM) was diluted to the concentration given in figures and tables at time t = 0, usually in 0.1 M Tris-HCl, pH 8.0 at 25°C. Subsequently, at short intervals, aliquots of the diluted enzyme solution were added to a cuvette and the activity was assayed as described above. During these experiments the specific activity of the enzyme gradually approached an equilibrium level of residual activity (Ar) that was retained for many hours. To determine this level of residual activity at several concentrations of enzyme, the enzyme stock was diluted to various concentrations for 200 min for the E206K-K296E double mutant enzyme or 330 min for the wild-type enzyme to acquire the final equilibrium level of activity (Ar).
Data treatment
According to Neet and Ainslie’s theory for hysteretic enzymes (Neet and Ainslie Jr. 1980), the decrease of enzymatic activity following dilution could be described by equation 1:
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 | ((2)) |
The KD value for each enzyme could also be determined from the saturation of remaining specific activity (Ar/[E]total) as a function of increasing protein concentration using the equation:
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The data were fitted to the equations using the BIOSOFT program UltraFit for Macintosh.
The free energy of dissociation of dimers to monomer subunits was obtained from the equation:
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Acknowledgments |
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. The financial support from the Danish National Science Council, the Danish National Research Foundation, and the Carlsberg Foundation is greatly appreciated. B.A.P. was supported by NIH Grant GM61087. The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.
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References |
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Argyrou, A., Washabaugh, M.W., and Pickart, C.M. 2000. Dihydroorotate dehydrogenase from Clostridium oroticum is a Class 1B enzyme and utilizes a concerted mechanism of catalysis. Biochemistry 39: 10373–10384.[CrossRef][Medline]
Björnberg, O., Rowland, P., Larsen, S., and Jensen, K.F. 1997. The active site of dihydroorotate dehydrogenase A from Lactococcus lactis investigated by chemical modification and mutagenesis. Biochemistry 36: 16197–16205.[CrossRef][Medline]
Björnberg, O., Grüner, A.C., Roepstorff, P., and Jensen, K.F. 1999. The activity of Escherichia coli dihydroorotate dehydrogenase is dependent on a conserved loop identified by sequence homology, mutagenesis and limited proteolysis. Biochemistry 28: 2899–2908.
Björnberg, O., Jordan, D.B., Palfey, B.A., and Jensen, K.F. 2001. Dihydrooxonate is a substrate of dihydroorotate dehydrogenase (DHOD) providing evidence for involvement of cysteine and serine residues in base catalysis. Arch. Biochem. Biophys. 391: 286–294.[CrossRef][Medline]
Deuschle, U., Kammerer, W., Gentz, R., and Bujard, H. 1986. Promoters of E. coli. A hierarchy of in vivo strength indicates alternate structures. EMBO J. 5: 2987–2994.[Medline]
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Marcinkeviciene, J., Tinney, L.M., Wang, K.H., Rogers, M.J., and Copeland, R.A. 1999. Dihydroorotate dehydrogenase B of Enterococcus faecalis. Characterization and insights into chemical mechanism. Biochemistry 38: 13129–13137.[CrossRef][Medline]
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Nielsen, F.S., Andersen, P.S., and Jensen, K.F. 1996a. The B-form of dihydroorotate dehydrogenase from Lactococcus lactis consists of two different subunits, encoded by the pyrDb and pyrK genes, and contains FMN, FAD, and [FeS] redox centres. J. Biol. Chem. 271: 29359–29365.
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, Ao = 28.5 ± 0.5 U/mg, Ar = 13.0 ± 0.5 U/mg, kobs = 0.033 ± 0.003 min-1), or 0.029 µM (
, Ao = 31.0 ± 0.7 U/mg, Ar = 4.9 ± 0.7 U/mg, kobs = 0.024 ± 0.002 min-1). (B) The E206K-K296E double mutant enzyme diluted to 9.8 µM (x, Ao = 29.3 ± 0.9 U/mg, Ar = 13.6 ± 0.3 U/mg, kobs = 0.146 ± 0.015 min-1), 0.98 µM (
, Ao = 26.3 ± 0.4 U/mg, Ar = 11.1 ± 0.3 U/mg, kobs = 0.026 ± 0.002 min-1). The curves are the best fits to the data points according to eq. 1
, E206K-K296E double mutant diluted in 0.1 M Tris-HCl, pH 8.0 containing 0.15 M NaCl. Following dilution at time t = 0, the samples were left to reach equilibrium for 200 min for the mutant DHODA and for 330 min for wild-type DHODA before assaying the remaining activity as described in Materials and Methods. The following parameters of the dissociation reaction were calculated according to eq. 3