Structure, catalytic mechanism, posttranslational lysine carbamylation, and inhibition of dihydropyrimidinases

Cheng-Yang Huanga,b,*

Abstract

Dihydropyrimidinase catalyzes the reversible hydrolytic ring opening of dihydrouracil and dihydrothymine to N-carbamoyl-b-alanine and N-carbamyl-b-aminoisobutyrate, respectively. Dihydropyrimidinase from microorganisms is normally known as hydantoinase because of its role as a biocatalyst in the synthesis of D- and L-amino acids for the industrial production of antibiotic precursors and its broad substrate specificity. Dihydropyrimidinase belongs to the cyclic amidohydrolase family, which also includes imidase, allantoinase, and dihydroorotase. Although these metal-dependent enzymes share low levels of amino acid sequence homology, they possess similar active site architectures and may use a similar mechanism for catalysis. By contrast, the five human dihydropyrimidinase-related proteins possess high amino acid sequence identity and are structurally homologous to dihydropyrimidinase, but they are neuronal proteins with no dihydropyrimidinase activity. In this chapter, we summarize and discuss current knowledge and the recent advances on the structure, catalytic mechanism, and inhibition of dihydropyrimidinase.

Summary

Pyrimidines are the key components in nucleic acids (Fig. 1) and are essential for the replication of genetic information in all biological systems (Del Cano-Ochoa, Moreno-Morcillo, & Ramon-Maiques, 2019; Evans & Guy, 2004; O’Donovan & Neuhard, 1970; Pardee & Yates, 1956). Pyrimidine is catabolized via oxidative and reductive pathways (Hayaishi & Kornberg, 1952; Vogels & Van der Drift, 1976). The reductive catabolism of pyrimidines proceeds in three sequential enzymatic steps via the enzymes dihydropyrimidine dehydrogenase (EC 1.3.1.2), dihydropyrimidinase (EC 3.5.2.2), and b-alanine synthase (EC 3.5.1.6) (Schnackerz & Dobritzsch, 2008). In the second step catalyzed by dihydropyrimidinase, dihydrouracil and dihydrothymine are converted to N-carbamoyl-b-alanine and N-carbamyl-b-aminoisobutyrate, respectively. This reaction is reversible, so dihydropyrimidinase also catalyzes the cyclization reaction in acidic environments.
Owing to its broad substrate specificity (Fig. 2), dihydropyrimidinase is also known as dihydropyrimidine hydrase (Wallach & Grisolia, 1957), 5,6-dihydropyrimidine amidohydrolase (Kautz & Schnackerz, 1989), hydantoinase (Eadie, Bernheim, & Bernheim, 1949), and imidase (Yang, Ramaswamy, & Jakoby, 1993). In addition to the physiological substrates dihydrouracil and dihydrothymine, dihydropyrimidinase catalyzes the hydrolytic cleavage of the linear and the heterocyclic imides (Yang et al., 1993). Phthalimide, 3,4-pyridine dicarboximide, 2,3-pyridine dicarboximide, 6-methyl dihydrouracil, 5-methyl dihydrouracil, succinimide, 2-phenylsuccinimide, glutarimide, adipimide, 1-methyl hydantoin, and 5-monosubstituted hydantoins are substrates of animal dihydropyrimidinase (Yang et al., 1993). The substrate spectrum of animal dihydropyrimidinase also includes cyclic carbonate (Yang, Ramaswamy, & Jakoby, 1998), maleimide (Huang & Yang, 2003), and N-iminylamide (Huang & Yang, 2005). Some sulfurcontaining cyclic imides can be hydrolyzed by imidase from Blastobacter sp. strain A17p-4 (Ogawa, Soong, Honda, & Shimizu, 1997).
Dihydropyrimidinase from microorganisms is normally known as hydantoinase because of its role as a biocatalyst (Fig. 3) in the synthesis of D- and Lamino acids for the industrial production of precursors for the semisynthesis of antibiotics, active peptides, hormones, and pesticides (Altenbuchner, Siemann-Herzberg, & Syldatk, 2001; Clemente-Jimenez, Martinez-Rodriguez, Rodriguez-Vico, & Heras-Vazquez, 2008; Schoemaker, Mink, & Wubbolts, 2003). Optically pure D-amino acids can be produced by the hydrolysis of D,L-5-monosubstituted hydantoins through the “hydantoinase process.” For example, D-p-hydroxyphenylglycine, a widely used intermediate for the synthesis of semi-synthetic antibiotics, can be produced from D,L-5-p-hydroxyphenylhydantoin by using D-hydantoinase in combination with D-carbamoylase. Alternatively, hydantoin racemase is used in this process for actively catalyzing the racemization of the unconverted L-hydantoin compound.
Almost all dihydropyrimidinases are tetramers (Fig. 4A). The global architecture of dihydropyrimidinase monomer consists of two domains, Structure, catalytic mechanism, posttranslational lysine carbamylation, and inhibition namely, a large domain with a classic (b/a)8-barrel structure core embedding the catalytic dimetal center (Fig. 4B) and a small b-sandwich domain. The binuclear metal center in dihydropyrimidinase consists of four His, one Asp, and one post-translational carbamylated Lys residues (Abendroth, Niefind, & Schomburg, 2002). Dihydropyrimidinase is a member of the cyclic amidohydrolase family (Gerlt & Babbitt, 2001; Kim & Kim, 1998b), which also includes dihydroorotase (Gojkovic et al., 2003; Grande-Garcia, Lallous, Diaz-Tejada, & Ramon-Maiques, 2014; Lee et al., 2007; Thoden, Phillips, Neal, Raushel, & Holden, 2001), allantoinase (Fig. 5) (Kim, Kim, Chung, Ahn, & Rhee, 2009), hydantoinase (Cheon et al., 2002), and imidase (Hung et al., 2010). Although the metal content of these metaldependent enzymes are somehow different, they possess similar active site architectures (Fig. 6) and catalyze the hydrolysis of the cyclic amide bond of each substrate in the metabolism of purines and pyrimidines (Tables 1e3). Given their involvement in the key reactions of nucleotide synthesis and degradation, some of these amidohydrolases are suggested as chemotherapeutic targets for anticancer (Huang, Lien, Chen, Lin, & Huang, 2020; Lee et al., 2018; Rabinovich et al., 2015), antimicrobial (Huang, 2015; Huang et al., 2020; Peng & Huang, 2014; Verrier et al., 2020), and antimalarial drug developments (Lee et al., 2007; Seymour, Lyons, Phillips, Rieckmann, & Christopherson, 1994).
Dihydropyrimidinase deficiency is an autosomal recessive disease characterized by a large accumulation of dihydrouracil and dihydrothymine in urine, blood, and cerebrospinal fluid (van Kuilenburg et al., 2010). Dihydropyrimidinase is a component in the chain for the biosynthesis of the neurotransmitter b-alanine, and its deficiency results in neurological abnormalities, including severe delay in speech development and white matter abnormalities (Henderson, Ward, Simmonds, Duley, & Davies, 1993; Tsuchiya et al., 2019; van Kuilenburg et al., 2007). Severe developmental delay is also found in infants with dihydropyrimidinase deficiency. Dihydropyrimidinase is the second enzyme in the catabolism of the anticancer drug 5-fluorouracil (5-FU), and 5-FU-associated toxicity is reported in asymptomatic patients with dihydropyrimidinase deficiency who underwent anticancer therapy (Sumi et al., 1998). Fig. 7 shows that 5-FU can bind to the active site of dihydropyrimidinase (Huang, Ning, & Huang, 2019). Many dihydropyrimidinase-like proteins such as collapsing response-mediator proteins, which highly resemble dihydropyrimidinase in terms of sequence and structure, are discovered (Table 4). Human dihydropyrimidinase shares an identity with dihydropyrimidinase-related proteins 1, 2, 3, 4, and 5 by 61%, 60%, 60%, 59%, and 58%, respectively and is higher than the Pseudomonas aeruginosa counterpart that shares amino acid residue identity with human dihydropyrimidinase by 51% (Fig. 8). Interestingly, these dihydropyrimidinase-related proteins are involved in neuronal development without the catalytic activity of dihydropyrimidinase (Wang & Strittmatter, 1997). The active site residues in dihydropyrimidinase are not conserved in human dihydropyrimidinase-related proteins, resulting in the absence of sites for metal binding and enzyme activity (Fig. 9).
Many studies on the biochemical properties and structural comparison with the mechanistic data of dihydropyrimidinase before 2009 have been thoroughly reviewed (Hsu, Lu, & Yang, 2010; Schnackerz & Dobritzsch, 2008). Over the last decade, investigations on the structure of dihydropyrimidinase from different species, as well as the inhibition modes, have provided considerable insights into the structureefunction relationships of dihydropyrimidinase. In this chapter, we review articles from 2010 to 2020 and summarize and discuss current knowledge and recent advances on the structure, catalytic mechanism, posttranslational lysine carbamylation, and inhibition of dihydropyrimidinases.

2. Structural mechanisms of Lys carbamylation indihydropyrimidinase

The metal center in the amidohydrolase superfamily is vital for enzymatic activities (Seibert & Raushel, 2005). Members of this family contain either a mononuclear metal center or a binuclear metal center (Seibert & Raushel, 2005). As a member of the amidohydrolase superfamily, dihydropyrimidinase/hydantoinase from different organisms contains one or two metal ions per subunit. The crystal structures of dihydropyrimidinase/ hydantoinase from Thermus sp. (Abendroth et al., 2002), Burkholderia pickettii (Xu et al., 2003), Bacillus stearothermophilus (Cheon et al., 2002), Saccharomyces kluyveri (Lohkamp, Andersen, Piskur, & Dobritzsch, 2006), Dictyostelium discoideum (Lohkamp et al., 2006), Sinorhizobium meliloti CECT4114 (Martinez-Rodriguez et al., 2010), and P. aeruginosa PAO1 (Tzeng, Huang, & Huang, 2016) are resolved. A carbamylated Lys located within the active site bridges the two metal ions (Fig. 4B). The crystal structures of human dihydropyrimidinase (Fig. 9A) and Bacillus sp. AR9 hydantoinase (Radha Kishan et al., 2005) are also determined, but the usual carbamylation on the active-site residue Lys does not happen. However, dihydropyrimidinase purified from bovine (Brooks, Jones, Kim, & Sander, 1983), rat (Kikugawa et al., 1994), pig (Huang & Yang, 2002), and fish liver (Huang & Yang, 2003) contains only a single divalent metal ion in the active site. Even in monometal-containing enzymes, the sites for dimetal binding in all these dihydropyrimidinases/hydantoinases are perfectly conserved.
The structures of apo-, monometal-, and dimetal-containing dihydropyrimidinase from Tetraodon nigroviridis (Fig. 10) reveal four steps in the assembly of the holoprotein with the carbamylated Lys and two metal ions (Hsieh et al., 2013). Structural and functional analyses indicate that only one metal (Zna) is sufficient for stabilizing carbamylation on the activesite residue Lys and for the maximal enzymatic activity of this fish dihydropyrimidinase (Hsieh et al., 2013). The structures of another monometallic dihydropyrimidinase from P. aeruginosa and the human dihydroorotase domain K1556A mutant further reveal that carbamylation on Lys is not required for Zna binding (Cheng, Huang, Lin, & Huang, 2018b). The presence or absence of Lys carbamylation does not affect Zna binding to the active site of dihydropyrimidinase. Thus, the stable carbamylation on the Lys in dihydropyrimidinase may be a Zna-mediated process. Although Lys carbamylation in dihydropyrimidinase does not need an additional enzyme to achieve covalent modification and is a spontaneous and reversible process, Zna may play a role in stabilizing the developing negative charge of carbamate (Fig. 10). When Lys is carbamylated, the negative charge of O facilitates additional metal coordination (Znb) for the regulation of specific enzymatic activities by controlling the conformations of two dynamic loops for the substrate entrance. This phenomenon may explain the ability of carboxylic acids to facilitate the assembly of the binuclear metal center only partially and temporarily (Ho, Huang, & Huang, 2013; Huang, Hsu, Chen, & Yang, 2009). Owing to different chemical properties between carbamate and carboxylate, only a resonance structure of carbamate can result in oxygen atoms forming negative charges (Fig. 11) that may promote di-metal assembly in dihydropyrimidinase. This phenomenon may also increase the nucleophilicity of the hydroxide in the active site of dihydropyrimidinase for catalysis (Kumar, Saxena, Sarma, & Radha Kishan, 2011).
The active site of dihydropyrimidinase may be grouped into six types: (I) a binuclear metal center with carbamylated Lys (Fig. 12A), (II) a binuclear metal center with uncarbamylated Lys (Fig. 12B), (III) a mononuclear metal center with carbamylated Lys (Fig. 12C), (IV) a mononuclear metal center with uncarbamylated Lys (Fig. 12D), (V) the site without any metal and carbamylated Lys (Fig. 12E), and (VI) the site without metal but with carbamylated Lys (Fig. 12F). With the analysis of enzyme activity as basis, the active site types I, II, and III are the catalytically active forms of dihydropyrimidinases found in dihydropyrimidinase/hydantoinase from Agrobacterium radiobacter (Huang et al., 2009), human (PDB entry 2VR2) and T. nigroviridis (Hsieh et al., 2013). At present, the active site type VI (no metal but with carbamylated Lys) is never found in any dihydropyrimidinase and in other cyclic amidohydrolases. Possibly, the spontaneous carbamylation on Lys must be stabilized by a metal ion within the active site of dihydropyrimidinase; otherwise, it cannot be maintained stably for a long time.

3. Molecular basis of dimer or tetramer formation: different contact residues

The global architecture of dihydropyrimidinase monomer consists of two domains: a large domain with a classic (b/a)8-barrel structure core embedding the catalytic dimetal center and a small b-sandwich domain. The length of a long C-terminal tail in dihydropyrimidinase varies among different species (Fig. 13AeF). Structural and functional analyses have indicated that all known dihydropyrimidinases/hydantoinases are tetramers except pseudomonal enzymes (Kim & Kim, 1998a; Niu, Zhang, Shi, & Yuan, 2007; Zhang, Niu, Shi, & Yuan, 2008). P. aeruginosa dihydropyrimidinase forms a dimer, rather than a tetramer, in the crystalline state and in the solution (Tzeng et al., 2016). The overall structure of each P. aeruginosa dihydropyrimidinase subunit consists of 17 a-helices and 19 b-sheets. The C-terminal tails of each subunit extend toward another monomer in a swapping-like manner (Fig. 13G), which may be involved in dihydropyrimidinase tetramerization (Lohkamp et al., 2006; Martinez-Rodriguez et al., 2010) and dimerization (Hsieh et al., 2013; Tzeng et al., 2016). Although P. aeruginosa dihydropyrimidinase has an extension C-terminal tail, P. aeruginosa dihydropyrimidinase forms a dimer (Fig. 13H and I). If any, contact is minimal, and many important and conserved residues are lacking in the dimeredimer interface of P. aeruginosa dihydropyrimidinase and are not enough to form a tetramer. However, the oligomerization of P. aeruginosa dihydropyrimidinase is a pH-dependent process (Cheng, Huang, Huang,

4. Catalytic mechanism: dynamic loops, tunnelbottleneck, and metal content of dihydropyrimidinases

Basing on their similar binuclear metal active site architecture and post-carbamylation, dihydropyrimidinase/hydantoinase and dihydroorotase share several enzymatic properties and may use a similar mechanism for catalysis (Gojkovic et al., 2003). For S. kluyveri dimetal-containing dihydropyrimidinase, two amino acids, Asn392 and Ser331, are directly involved in substrate and product binding. The binding of this enzyme to the reaction product proceeds in a manner similar to that of an enzyme and a substrate (Schnackerz & Dobritzsch, 2008). Notably, no solvent molecules are involved in ligand binding. A narrow channel probably used for substrate entry is found from the protein surface on the active site of S. kluyveri dimetal-containing dihydropyrimidinase. Tyr172 located within a dynamic loop in this enzyme may play an essential role in the stabilization of the tetrahedral transition state during hydrolysis of the substrate, collapse of the transition state, product formation, and product release (Cheon et al., 2002; Lohkamp et al., 2006).
Two dynamic loops (segments Ala69eArg74 and Met158eMet165) and the C-terminal tail are important for the substrate entrance of T. nigroviridis dihydropyrimidinase. Tyr160 that structurally corresponds to the essential Tyr172 residue in S. kluyveri dihydropyrimidinase is also found in T. nigroviridis dihydropyrimidinase (Hsieh et al., 2013). Three stages are proposed on the basis of the structures of the apo- (the open form), monometal(the intermediate stage), and dimetal-containing dihydropyrimidinases (the closed form) from T. nigroviridis. The cross-section bottlenecks of the active site pocket and the conformations of dynamic loops among the three stages of this enzyme are significantly different. The tunnel bottlenecks in the open and closed forms of T. nigroviridis dihydropyrimidinase are approximately 10 and 3 Å, respectively. Given that the tunnel bottleneck in the closed form (dimetallic enzyme) is narrower than the average size of substrates (6 Å), the bottleneck size may indicate a substrate entrance. However, the dynamic process for substrate diffusion and different catalytic mechanisms make whether this assumption is applicable to cases of dimetallic dihydropyrimidinase unclear (Fig. 15). For example, the bottleneck sizes of the tunnel in the monometallic (Cheng et al., 2018b) and dimetallic P. aeruginosa dihydropyrimidinase (Tzeng et al., 2016) are 4.5 and 4.0 Å, respectively. Both values suggest a closed form, but the dimetallic P. aeruginosa dihydropyrimidinase is catalytically active.
The active form of animal dihydropyrimidinases is a monometallic enzyme (Brooks et al., 1983; Huang & Yang, 2002, 2003; Kikugawa et al., 1994). In addition to structural analysis, the linear titration curve for apo-form enzyme reactivation with zinc ions indicates that only one metal ion is required for the active T. nigroviridis dihydropyrimidinase (Fig. 16A) (Hsieh et al., 2013). This phenomenon may explain why the metal ion becomes an inhibitor as soon as the first metal binding site is occupied (Hsieh et al., 2013). Binding of the second zinc ion to T. nigroviridis dihydropyrimidinase induces a conformational change on the bottleneck of the active site tunnel, thereby blocking the site for substrate binding as also seen in thermolysin (Holland, Hausrath, Juers, & Matthews, 1995). The structural and biochemical analyses demonstrate that the catalytic mechanism of T. nigroviridis dihydropyrimidinase somehow differs from that of the dimetallic dihydropyrimidinases and dihydroorotases (Fig. 16B) (Gojkovic et al., 2003) and provide a distinctive mechanistic view currently held in the field. The human dihydropyrimidinase is a dimetal-containing enzyme with uncarbamylated Lys, but whether this form is catalytically active or is a (second) metal ion-inhibited form remains unclear (Fig. 9A). Perhaps, the titration curve for reactivating the apo-form enzyme with zinc ions should be plotted to determine the active form of human dihydropyrimidinase. A metal titration curve showing a nonlinear increase in the activity of apo-form enzyme suggests that a kinetic model in which only dihydropyrimidinase with a fully assembled binuclear metal center (M1M2-dihydropyrimidinase) is catalytically active. The first metal (M1) is preferentially bound to the binuclear center (Fig. 16B), as in the case of A. radiobacter hydantoinase (Huang et al., 2009).
Although human dihydropyrimidinase and dihydropyrimidinase-related proteins share a high level of sequence homology (with 58%e61% identity and 73%e77% similarity), dihydropyrimidinase-related proteins are not the enzymes responsible for dihydrouracil and dihydrothymine hydrolysis. The active site metal-binding residues in dihydropyrimidinase are not conserved in human dihydropyrimidinase-related proteins, leading to the absence of a catalytic activity (Fig. 9). However, the substrate-binding residue (Tyr164), as well as the dynamic loop (segment Met162eMet169) for substrate diffusion in human dihydropyrimidinase, is somehow similar in these neuroproteins. Whether the big cavity in these dihydropyrimidinase-related proteins is responsible for some unknown molecule accommodation and interaction is not clear (Fig. 9BeF). For example, the essential Tyr residue in human dihydropyrimidinase is almost conserved in dihydropyrimidinase-related proteins (Fig. 8). Whether these neuroproteins can bind dihydrouracil and the reaction product (e.g., N-carbamoyl-b-alanine) remains unknown. b-alanine is a small molecule neurotransmitter widely distributed in the central nervous system and the reductive degradation of uracil in mammals is the sole source of b-alanine. Whether the empty cavity in these dihydropyrimidinase-related proteins is associated with binding or transporting dihydrouracil via the dynamic loop should be further studied via structural and functional identification.

5. Competitive inhibitors of dihydropyrimidinase

Considering that dihydropyrimidinase is a component in the chain of pyrimidine catabolism required for metabolizing the DNA base, blocking its activity may be useful to limit bacterial and cancer cell growth and survival. Substrate analogs for any enzyme are usually potential inhibitors, but this common rule is not applicable to dihydropyrimidinase. Possibly due to the broad substrate specificity of dihydropyrimidinase, inhibitors that target its active site are not enough to strongly suppress its activity. Although some chelators, such as 8-hydroxyquinoline-5-sulfonic acid (Brooks et al., 1983), inhibit dihydropyrimidinase, they are harmful to humans. Reaction products, such as N-carbamyl-b-alanine (with Ki value of 0.68 mM), are inhibitors of dihydropyrimidinase (Kikugawa et al., 1994). Product analogs glutarate monoamide and 4-ureidobutyrate with Ki values of 0.21 and 0.99 mM, respectively, inhibit dihydropyrimidinase (Kikugawa et al., 1994). High concentrations of dihydroorotate, 5-hydantoin acetic acid, and acetohydroxamate (within the millimolar range) are required to greatly inhibit dihydropyrimidinase (Huang, 2015). Thus, substrate/product analogs do not work as potent inhibitors on dihydropyrimidinase.
Given the poor inhibition ability of substrate analogs, screening new dihydropyrimidinase inhibitors from natural products may be a good strategy. Plumbagin isolated from the extract of the carnivorous plant Nepenthes miranda is an inhibitor of P. aeruginosa dihydropyrimidinase with a half maximal inhibitory concentration (IC50) value of 58 mM (Huang et al., 2020). The flavonoid dihydromyricetin significantly inhibits human (Basbous et al., 2020) and P. aeruginosa dihydropyrimidinase activity (Huang, 2015) with IC50 values of approximately 6 and 48 mM, respectively. These naturally occurring compounds seem to inhibit dihydropyrimidinase through similar but different modes by competing with dihydrouracil for the active sites of dihydropyrimidinase. According to the docking models of P. aeruginosa dihydropyrimidinase (Fig. 17), dihydromyricetin interacts with Ile95, Ser289, and Asp316 (Huang, 2015), whereas plumbagin interacts with Tyr155, Lys156, Val190, Gln194, Arg212, and Asn337 (Huang et al., 2020). Residues Tyr155, Ser289, and Asn337 are crucial for substrate binding, and Asp316 has essential dual roles in metal Zna binding and in catalysis in P. aeruginosa dihydropyrimidinase. Thus, these compounds partially occupy and block the active site of dihydropyrimidinase for inhibition. In addition, Tyr155 is the most important residue within the dynamic loop involved in stabilizing the intermediate state during catalysis (Hsieh et al., 2013). Further research can focus on designing compounds that target the dynamic loops and catalytic sites in dihydropyrimidinase for inhibitor optimization and drug development (Huang et al., 2020).
Over the last several years, enzymes involved in pyrimidine nucleotide metabolism as targets for chemotherapy have been intensively studied. Many pyrimidine analogs, such as azidothymidine and 5-FU, have been used as antiviral and anticancer chemotherapeutic drugs. Regulation of dihydropyrimidinase activity may induce different effects on living cells. Recently, crystal structures for monometallic dihydropyrimidinase, as well as the complex with 5-FU, are determined. The elucidation of different crystal structures of dihydropyrimidinase, together with biochemical studies, will be helpful in providing new knowledge of their structureefunction relationships. Some natural products (but not substrate analogs) as inhibitors on dihydropyrimidinase are discovered. Designing and finding potent inhibitors remain a challenge. Further studies are necessary for inhibition and drug optimization on dihydropyrimidinase and finding promising drugs for suppressing pyrimidine metabolism.

References

Abendroth, J., Niefind, K., & Schomburg, D. (2002). X-ray structure of a dihydropyrimidinase from Thermus sp. at 1.3 A resolution. Journal of Molecular Biology, 320(1), 143e156.
Altenbuchner, J., Siemann-Herzberg, M., & Syldatk, C. (2001). Hydantoinases and related enzymes as biocatalysts for the synthesis of unnatural chiral amino acids. Current Opinion in Biotechnology, 12(6), 559e563.
Basbous, J., Aze, A., Chaloin, L., Lebdy, R., Hodroj, D., Ribeyre, C., et al. (2020). Dihydropyrimidinase protects from DNA replication stress caused by cytotoxic metabolites. Nucleic Acids Research, 48(4), 1886e1904.
Brooks, K. P., Jones, E. A., Kim, B. D., & Sander, E. G. (1983). Bovine liver dihydropyrimidine amidohydrolase: Purification, properties, and characterization as a zinc metalloenzyme. Archives of Biochemistry and Biophysics, 226(2), 469e483.
Cheng, J. H., Huang, C. C., Huang, Y. H., & Huang, C. Y. (2018a). Structural basis for pHdependent oligomerization of dihydropyrimidinase from Pseudomonas aeruginosa PAO1. Bioinorganic Chemistry and Applications, 2018, 9564391.
Cheng, J. H., Huang, Y. H., Lin, J. J., & Huang, C. Y. (2018b). Crystal structures of monometallic dihydropyrimidinase and the human dihydroorotase domain K1556A mutant reveal no lysine carbamylation within the active site. Biochemical and Biophysical Research Communications, 505(2), 439e444.
Cheon, Y. H., Kim, H. S., Han, K. H., Abendroth, J., Niefind, K., Schomburg, D., et al. (2002). Crystal structure of D-hydantoinase from Bacillus stearothermophilus: Insight into the stereochemistry of enantioselectivity. Biochemistry, 41(30), 9410e9417.
Clemente-Jimenez, J. M., Martinez-Rodriguez, S., Rodriguez-Vico, F., & HerasVazquez, F. J. (2008). Optically pure alpha-amino acids production by the “Hydantoinase Process”. Recent Patents on Biotechnology, 2(1), 35e46.
Del Cano-Ochoa, F., Moreno-Morcillo, M., & Ramon-Maiques, S. (2019). CAD, A multienzymatic protein at the head of de Novo pyrimidine biosynthesis. Subcellular Biochemistry, 93, 505e538.
Eadie, G. S., Bernheim, F., & Bernheim, M. L. (1949). The partial purification and properties of animal and plant hydantoinases. Journal of Biological Chemistry, 181(2), 449e458.
Evans, D. R., & Guy, H. I. (2004). Mammalian pyrimidine biosynthesis: Fresh insights into an ancient pathway. Journal of Biological Chemistry, 279(32), 33035e33038.
Gerlt, J. A., & Babbitt, P. C. (2001). Divergent evolution of enzymatic function: Mechanistically diverse superfamilies and functionally distinct suprafamilies. Annual Review of Biochemistry, 70, 209e246.
Gojkovic, Z., Rislund, L., Andersen, B., Sandrini, M. P., Cook, P. F., Schnackerz, K. D., et al. (2003). Dihydropyrimidine amidohydrolases and dihydroorotases share the same origin and several enzymatic properties. Nucleic Acids Research, 31(6), 1683e1692.
Grande-Garcia, A., Lallous, N., Diaz-Tejada, C., & Ramon-Maiques, S. (2014). Structure, functional characterization, and evolution of the dihydroorotase domain of human CAD. Structure, 22(2), 185e198.
Hayaishi, O., & Kornberg, A. (1952). Metabolism of cytosine, thymine, uracil, and barbituric acid by bacterial enzymes. Journal of Biological Chemistry, 197(2), 717e732.
Henderson, M. J., Ward, K., Simmonds, H. A., Duley, J. A., & Davies, P. M. (1993). Dihydropyrimidinase deficiency presenting in infancy with severe developmental delay. Journal of Inherited Metabolic Disease, 16(3), 574e576.
Ho, Y. Y., Huang, Y. H., & Huang, C. Y. (2013). Chemical rescue of the post-translationally carboxylated lysine mutant of allantoinase and dihydroorotase by metal ions and shortchain carboxylic acids. Amino Acids, 44(4), 1181e1191.
Holland, D. R., Hausrath, A. C., Juers, D., & Matthews, B. W. (1995). Structural analysis of zinc substitutions in the active site of thermolysin. Protein Science, 4(10), 1955e1965.
Hsieh, Y. C., Chen, M. C., Hsu, C. C., Chan, S. I., Yang, Y. S., & Chen, C. J. (2013). Crystal structures of vertebrate dihydropyrimidinase and complexes from Tetraodon nigroviridis with lysine carbamylation: Metal and structural requirements for post-translational modification and function. Journal of Biological Chemistry, 288(42), 30645e30658.
Hsu, C. C., Lu, L. Y., & Yang, Y. S. (2010). From sequence and structure of sulfotransferases and dihydropyrimidinases to an understanding of their mechanisms of action and function. Expert Opinion on Drug Metabolism and Toxicology, 6(5), 591e601.
Huang, C. Y. (2015). Inhibition of a putative dihydropyrimidinase from Pseudomonas aeruginosa PAO1 by flavonoids and substrates of cyclic amidohydrolases. PLoS One, 10(5), e0127634.
Huang, C. Y., Hsu, C. C., Chen, M. C., & Yang, Y. S. (2009). Effect of metal binding and posttranslational lysine carboxylation on the activity of recombinant hydantoinase. Journal of Biological Inorganic Chemistry, 14(1), 111e121.
Huang, Y. H., Lien, Y., Chen, J. H., Lin, E. S., & Huang, C. Y. (2020). Identification and characterization of dihydropyrimidinase inhibited by plumbagin isolated from Nepenthes miranda extract. Biochimie, 171e172, 124e135.
Huang, Y. H., Ning, Z. J., & Huang, C. Y. (2019). Crystal structure of dihydropyrimidinase in complex with anticancer drug 5-fluorouracil. Biochemical and Biophysical Research Communications, 519(1), 160e165.
Huang, C. Y., & Yang, Y. S. (2002). The role of metal on imide hydrolysis: Metal content and pH profiles of metal ion-replaced mammalian imidase. Biochemical and Biophysical Research Communications, 297(4), 1027e1032.
Huang, C. Y., & Yang, Y. S. (2003). A novel cold-adapted imidase from fish Oreochromis niloticus that catalyzes hydrolysis of maleimide. Biochemical and Biophysical Research Communications, 312(2), 467e472.
Huang, C. Y., & Yang, Y. S. (2005). Discovery of a novel N-iminylamidase activity: Substrate specificity, chemicoselectivity and catalytic mechanism. Protein Expression and Purification, 40(1), 203e211.
Hung, C. L., Lee, C., Lin, C. Y., Chang, C. H., Chung, Y. C., & Yi Tang, C. (2010). Feature amplified voting algorithm for functional analysis of protein superfamily. BMC Genomics, 11(Suppl. 3), S14.
Kautz, J., & Schnackerz, K. D. (1989). Purification and properties of 5,6-dihydropyrimidine amidohydrolase from calf liver. European Journal of Biochemistry, 181(2), 431e435.
Kikugawa, M., Kaneko, M., Fujimoto-Sakata, S., Maeda, M., Kawasaki, K., Takagi, T., et al. (1994). Purification, characterization and inhibition of dihydropyrimidinase from rat liver. European Journal of Biochemistry, 219(1e2), 393e399.
Kim, G. J., & Kim, H. S. (1998a). C-terminal regions of D-hydantoinases are nonessential for catalysis, but affect the oligomeric structure. Biochemical and Biophysical Research Communications, 243(1), 96e100.
Kim, G. J., & Kim, H. S. (1998b). Identification of the structural similarity in the functionally related amidohydrolases acting on the cyclic amide ring. Biochemical Journal, 330(Pt 1), 295e302.
Kim, K., Kim, M. I., Chung, J., Ahn, J. H., & Rhee, S. (2009). Crystal structure of metaldependent allantoinase from Escherichia coli. Journal of Molecular Biology, 387(5), 1067e 1074. van Kuilenburg, A. B., Dobritzsch, D., Meijer, J., Meinsma, R., Benoist, J. F., Assmann, B., et al. (2010). Dihydropyrimidinase deficiency: Phenotype, genotype and structural consequences in 17 patients. Biochimica et Biophysica Acta, 1802(7e8), 639e648.
van Kuilenburg, A. B., Meijer, J., Dobritzsch, D., Meinsma, R., Duran, M., Lohkamp, B., et al. (2007). Clinical, biochemical and genetic findings in two siblings with a dihydropyrimidinase deficiency. Molecular Genetics and Metabolism, 91(2), 157e164.
Kumar, V., Saxena, N., Sarma, M., & Radha Kishan, K. V. (2011). Carboxylated lysine is required for higher activities in Hydantoinases. Protein and Peptide Letters, 18(7), 663e 669.
Lee, J. S., Adler, L., Karathia, H., Carmel, N., Rabinovich, S., Auslander, N., et al. (2018). Urea cycle dysregulation generates clinically relevant genomic and biochemical signatures. Cell, 174(6), 1559e1570.e22.
Lee, M., Chan, C. W., Graham, S. C., Christopherson, R. I., Guss, J. M., & Maher, M. J. (2007). Structures of ligand-free and inhibitor complexes of dihydroorotase from Escherichia coli: Implications for loop movement in inhibitor design. Journal of Molecular Biology, 370(5), 812e825.
Lohkamp, B., Andersen, B., Piskur, J., & Dobritzsch, D. (2006). The crystal structures of dihydropyrimidinases reaffirm the close relationship between cyclic amidohydrolases and explain their substrate specificity. Journal of Biological Chemistry, 281(19), 13762e 13776.
Martinez-Rodriguez, S., Martinez-Gomez, A. I., Clemente-Jimenez, J. M., RodriguezVico, F., Garcia-Ruiz, J. M., Las Heras-Vazquez, F. J., et al. (2010). Structure of dihydropyrimidinase from Sinorhizobium meliloti CECT4114: New features in an amidohydrolase family member. Journal of Structural Biology, 169(2), 200e208.
Niu, L., Zhang, X., Shi, Y., & Yuan, J. (2007). Subunit dissociation and stability alteration of D hydantoinase deleted at the terminal amino acid residue. Biotechnology Letters, 29(2), 303e308.
O’Donovan, G. A., & Neuhard, J. (1970). Pyrimidine metabolism in microorganisms. Bacteriological Reviews, 34(3), 278e343.
Ogawa, J., Soong, C. L., Honda, M., & Shimizu, S. (1997). Imidase, a dihydropyrimidinaselike enzyme involved in the metabolism of cyclic imides. European Journal of Biochemistry, 243(1e2), 322e327.
Pardee, A. B., & Yates, R. A. (1956). Pyrimidine biosynthesis in Escherichia coli. Journal of Biological Chemistry, 221(2), 743e756.
Peng, W. F., & Huang, C. Y. (2014). Allantoinase and dihydroorotase binding and inhibition by flavonols and the substrates of cyclic amidohydrolases. Biochimie, 101, 113e122.
Rabinovich, S., Adler, L., Yizhak, K., Sarver, A., Silberman, A., Agron, S., et al. (2015). Diversion of aspartate in ASS1-deficient tumours fosters de novo pyrimidine synthesis. Nature, 527(7578), 379e383.
Radha Kishan, K. V., Vohra, R. M., Ganesan, K., Agrawal, V., Sharma, V. M., & Sharma, R. (2005). Molecular structure of D-hydantoinase from Bacillus sp. AR9: Evidence for mercury inhibition. Journal of Molecular Biology, 347, 95e105.
Schnackerz, K. D., & Dobritzsch, D. (2008). Amidohydrolases of the reductive pyrimidine catabolic pathway purification, characterization, structure, reaction mechanisms and enzyme deficiency. Biochimica et Biophysica Acta, 1784(3), 431e444.
Schoemaker, H. E., Mink, D., & Wubbolts, M. G. (2003). Dispelling the myths–biocatalysis in industrial synthesis. Science, 299(5613), 1694e1697.
Seibert, C. M., & Raushel, F. M. (2005). Structural and catalytic diversity within the amidohydrolase superfamily. Biochemistry, 44(17), 6383e6391.
Seymour, K. K., Lyons, S. D., Phillips, L., Rieckmann, K. H., & Christopherson, R. I. (1994). Cytotoxic effects of inhibitors of de novo pyrimidine biosynthesis upon Plasmodium falciparum. Biochemistry, 33(17), 5268e5274.
Sumi, S., Imaeda, M., Kidouchi, K., Ohba, S., Hamajima, N., Kodama, K., et al. (1998). Population and family studies of dihydropyrimidinuria: Prevalence, inheritance mode, and risk of fluorouracil toxicity. American Journal of Medical Genetics, 78(4), 336e340.
Thoden, J. B., Phillips, G. N., Jr., Neal, T. M., Raushel, F. M., & Holden, H. M. (2001). Molecular structure of dihydroorotase: A paradigm for catalysis through the use of a binuclear metal center. Biochemistry, 40(24), 6989e6997.
Tsuchiya, H., Akiyama, T., Kuhara, T., Nakajima, Y., Ohse, M., Kurahashi, H., et al. (2019). A case of dihydropyrimidinase deficiency incidentally detected by urine metabolome analysis. Brain & Development, 41(3), 280e284.
Tzeng, C. T., Huang, Y. H., & Huang, C. Y. (2016). Crystal structure of dihydropyrimidinase from Pseudomonas aeruginosa PAO1: Insights into the molecular basis of formation of a dimer. Biochemical and Biophysical Research Communications, 478(3), 1449e1455.
Verrier, E. R., Weiss, A., Bach, C., Heydmann, L., Turon-Lagot, V., Kopp, A., et al. (2020). Combined small molecule and loss-of-function screen uncovers estrogen receptor alpha and CAD as host factors for HDV infection and antiviral targets. Gut, 69(1), 158e167.
Vogels, G. D., & Van der Drift, C. (1976). Degradation of purines and pyrimidines by microorganisms. Bacteriological Reviews, 40(2), 403e468.
Wallach, D. P., & Grisolia, S. (1957). The purification and properties of hydropyrimidine hydrase. Journal of Biological Chemistry, 226(1), 277e288.
Wang, L. H., & Strittmatter, S. M. (1997). Brain CRMP forms heterotetramers similar to liver dihydropyrimidinase. Journal of Neurochemistry, 69(6), 2261e2269.
Xu, Z., Liu, Y., Yang, Y., Jiang, W., Arnold, E., & Ding, J. (2003). Crystal structure of Dhydantoinase from Burkholderia pickettii at a resolution of 2.7 angstroms: Insights into the molecular basis of enzyme thermostability. Journal of Bacteriology, 185(14), 4038e 4049.
Yang, Y. S., Ramaswamy, S., & Jakoby, W. B. (1993). Rat liver imidase. Journal of Biological Chemistry, 268(15), 10870e10875.
Yang, Y. L., Ramaswamy, S. G., & Jakoby, W. B. (1998). Enzymatic hydrolysis of organic cyclic carbonates. Journal of Biological Chemistry, 273(14), 7814e7817.
Zhang, X. Y., Niu, L. X., Shi, Y. W., & Yuan, J. M. (2008). The flexibility of the non-conservative region at the C terminus of D-hydantoinase from Pseudomonas putida YZ-26 is extremely limited. Applied Biochemistry and Biotechnology, 144(3), 237e247.