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Flavoprotein quinone reductases regenerate quinols which serve metabolic and antioxidant roles. These enzymes catalyze the two-electron oxidation of substrates and the subsequent two electron reduction of quinones. Despite the net two electron transfer between substrates, the binding mode of quinones is typically end-on to the flavin, rather than stacked, dictating that the oxidative half reaction cannot proceed via hydride transfer and must instead occur by two successive single electron transfers. Here we present a review of six of the most well-studied flavoprotein quinone reductases to establish a framework for discussing this positional orientation for the quinone oxidant. There are two non-mutually exclusive rationalizations for this binding mode where the flavin isoalloxazine acts as a redox partition. The first is that energetics of the single electron transfer pathway create a kinetic barrier to the reverse reaction, trapping electrons in the quinone pool and countering the high ratio of quinol to quinone present in the membrane. The second is that the end-on binding allows the enzymes to utilize different binding sites for cytosolic and membrane associated substrates, avoiding the need to desorb substrates. These effects may be additive and serve to funnel electrons into the quinone pool as efficiently as possible. 

Human ferroptosis suppressor protein 1 (HsFSP1) is an NAD(P)H:quinone oxidoreductase with broad substrate specificity that has been widely implicated in aiding malignant neoplastic cell survival. FSP1 is myristoylated and associated with membranes, where it regenerates the reduced forms of quinones using electrons from NADPH. The quinol products intercept reactive oxygen species and ameliorate lipid peroxidation, preventing ferroptosis, a form of regulated cell death. While FSP1 enzymes have been reported to have 6-OH-FAD as an active cofactor, aerobic titration of the enzyme with NADPH in the presence and absence of ubiquinone (UQ) reveals that this is more likely an artifact and that the native form of HsFSP1 has unmodified FAD as the cofactor. Moreover, HsFSP1 suppresses the reaction of the reduced FAD with molecular oxygen three-fold which, from a kinetic standpoint, severely limits the opportunity for cofactor modification. The isolated form of the enzyme has NADP+ bound and the rate of release of this product limits the observed rate of reduction by NAD(P)H molecules. The reduction of substrate quinones occurs rapidly (≥2000 s–1), dictating that the rate of turnover is wholly defined by the rate of release of NADP+ from the HsFSP1·NADP+ complex. Given that HsFSP1 does not distinguish ubiquinone from ubiquinol by significant differences in binding affinity, this pronounced catalytic commitment to quinone reduction serves to overcome presumed kinetic limitations imposed by the abundance of ubiquinol relative to ubiquinone in the membrane. This characteristic also maintains the enzyme ostensibly fully in the oxidized state under turnover conditions, preventing significant futile reduction of dioxygen. 

Thioredoxin/glutathione reductase from Schistosoma mansoni (SmTGR) is a multifunctional enzyme that catalyzes the reduction of glutathione (GSSG) and thioredoxin, as well as the deglutathionylation of peptide and non-peptide substrates. SmTGR structurally resembles known glutathione reductases (GR) and thioredoxin reductases (TrxR) but with an appended N-terminal domain that has a typical glutaredoxin (Grx) fold. Despite structural homology with known GRs, the site of GSSG reduction has frequently been reported as the Grx domain, based primarily on aerobic, steady-state kinetic measurements and x-ray crystallography. Here, we present an anaerobic characterization of a series of variant SmTGRs to establish the site of GSSG reduction as the cysteine pair most proximal to the FAD, Cys154/Cys159, equivalent to the site of GSSG reduction in GRs. Anaerobic steady-state analysis of U597C, U597S, U597C + C31S, and I592STOP SmTGR demonstrate that the Grx domain is not involved in the catalytic reduction of GSSG, as redox silencing of the C-terminus results in no modulation of the observed turnover number (∼0.025 s−1) and redox silencing of the Grx domain results in an increased observed turnover number (∼0.08 s−1). Transient-state single turnover analysis of these variants corroborates this, as the slowest rate observed titrates hyperbolically with GSSG concentration and approaches a limit that coincides with the respective steady-state turnover number for each variant. Numerical integration fitting of the transient state data can only account for the observed trends when competitive binding of the C-terminus is included, indicating that the partitioning of electrons to either substrate occurs at the Cys154/Cys159 disulfide rather than the previously proposed Cys596/Sec597 sulfide/selenide. Paradoxically, truncating the C-terminus at Ile592 results in a loss of GR activity, indicating a crucial non-redox role for the C-terminus. 

Dihydropyrimidine dehydrogenase (DPD) is an enzyme that uses an elaborate architecture to catalyze a simple net reaction: the reduction of the vinylic bond of uracil and thymine. Known DPDs have two active sites separated by approximately 60 Å. One active site has an FAD cofactor and binds NAD(P) and the other has an FMN cofactor and binds pyrimidines. The intervening distance is spanned by four Fe4S4 centers that act as an electron conduit. Recent advancements with porcine DPD have revealed unexpected chemical sequences where the enzyme undergoes reductive activation by transferring two electrons from NADPH to the FMN via the FAD such that the active form has the cofactor set FAD•4(Fe4S4)•FMNH2. Here we describe the first comprehensive kinetic investigation of a bacterial form of DPD. Using primarily transient state methods, DPD from E. coli (EcDPD) was shown to have a similar mechanism to that observed with the mammalian form in that EcDPD is observed to undergo reductive activation before pyrimidine reduction and displays half-of-sites activity. However, two distinct aspects of the EcDPD reaction relative to the mammalian enzyme were observed that relate to the effector roles for substrates: (i) the enzyme will rapidly take up electrons from NADH, reducing a flavin in the absence of pyrimidine substrate, and (ii) the activated form of the enzyme can become fully oxidized by transferring electrons to pyrimidine substrates in the absence of NADH. 

Human ornithine aminotransferase (hOAT), a pyridoxal 5'-phosphate (PLP)-dependent enzyme, has been shown to play an essential role in the metabolic reprogramming and progression of hepatocellular carcinoma (HCC). HCC accounts for approximately 75% of primary liver cancers and is within the top three causes of cancer death worldwide. As a result of treatment limitations, the overall 5-year survival rate for all patients with HCC is under 20%. The prevalence of HCC necessitates continued development of novel and effective treatment methods. In recent years, the therapeutic potential of selective inactivation of hOAT has been demonstrated for the treatment of HCC. Inspired by previous increased selectivity for hOAT by the expansion of the cyclopentene ring scaffold to a cyclohexene, we designed, synthesized, and evaluated a series of novel fluorinated cyclohexene analogues and identified (R)-3-amino-5,5-difluorocyclohex-1-ene-1-carboxylic acid as a time-dependent inhibitor of hOAT. Structural and mechanistic studies have elucidated the mechanism of inactivation of hOAT by 5, resulting in a PLP-inactivator adduct tightly bound to the active site of the enzyme. Intact protein mass spectrometry, 19F NMR spectroscopy, transient state kinetic studies, and X-ray crystallography were used to determine the structure of the final adduct and elucidate the mechanisms of inactivation. Interestingly, despite the highly electrophilic intermediate species conferred by fluorine and structural evidence of solvent accessibility in the hOAT active site, Lys292 and water did not participate in nucleophilic addition during the inactivation mechanism of hOAT by 5. Instead, rapid aromatization to yield the final adduct was favored. 

Thioredoxin/glutathione reductase from Schistosoma mansoni (SmTGR) catalyzes the reduction of both oxidized thioredoxin and glutathione with electrons from reduced nicotinamide adenine dinucleotide phosphate (NADPH). SmTGR is a drug target for the treatment of Schistosomiasis, an infection caused by Schistosoma platyhelminths residing in the blood vessels of the host. Schistosoma spp. are reliant on TGR enzymes as they lack catalase and so use reduced thioredoxin and glutathione to regenerate peroxiredoxins consumed in the detoxification of reactive oxygen species. SmTGR is a flavin adenine dinucleotide (FAD)-dependent enzyme, and we have used the flavin as a spectrophotometric reporter to observe the movement of electrons within the enzyme. The data show that NADPH fractionally reduces the active site flavin with an observed rate constant estimated in this study to be ∼3000 s-1. The flavin then reoxidizes by passing electrons at a similar rate to the proximal Cys159-Cys154 disulfide pair. The dissociation of NADP+ occurs with a rate of ∼180 s-1, which induces the deprotonation of Cys159, and this coincides with the accumulation of an intense FAD-thiolate charge transfer band. It is proposed that the electrons then pass to the Cys596-Cys597 disulfide pair of the associated subunit in the dimer with a net rate constant of ∼2 s-1. (Note: Cys597 is Sec597 in wild-type (WT) SmTGR.) From this position, the electrons can be passed to oxidized thioredoxin or further into the protein to reduce the Cys28-Cys31 disulfide pair of the originating subunit of the dimer. From the Cys28-Cys31 center, electrons can then pass to oxidized glutathione that has a binding site directly adjacent.